Ligand–receptor interactions that are reinforced by mechanical stress, so-called catch-bonds, play a major role in cell–cell adhesion. They critically contribute to widespread urinary tract infections by pathogenic Escherichia coli strains. These pathogens attach to host epithelia via the adhesin FimH, a two-domain protein at the tip of type I pili recognizing terminal mannoses on epithelial glycoproteins. Here we establish peptide-complemented FimH as a model system for fimbrial FimH function. We reveal a three-state mechanism of FimH catch-bond formation based on crystal structures of all states, kinetic analysis of ligand interaction and molecular dynamics simulations. In the absence of tensile force, the FimH pilin domain allosterically accelerates spontaneous ligand dissociation from the FimH lectin domain by 100,000-fold, resulting in weak affinity. Separation of the FimH domains under stress abolishes allosteric interplay and increases the affinity of the lectin domain. Cell tracking demonstrates that rapid ligand dissociation from FimH supports motility of piliated E. coli on mannosylated surfaces in the absence of shear force.
Cell–cell adhesion often occurs under dynamically varying conditions and mechanical stress. In many cell–cell adhesion systems, the lifetime of adhesin–receptor complexes is increased under tensile mechanical force via ‘catch-bonds’, which permit capture or retention of cells under flow conditions while still allowing for release under reduced mechanical force. Catch-bond interactions are prominent in vascular systems and are formed, for example, by selectins for leukocyte recruitment1,2, by cadherins controlling tissue integrity3,4 in the epithelial adhesion of cancer cells5 and by the interactions between T-cell receptors (TCRs) and peptide-bound major histocompatibility complexes (MHC) on antigen-presenting cells6,7. Catch-bonds also play a major role in bacterial adhesion and infection by uropathogenic Escherichia coli strains, which are responsible for the vast majority of urinary tract infections (UTIs) in humans8. A first critical step in the establishment of infection is bacterial adhesion to urothelial cells under flow conditions, which is mediated by 0.1−2 μm long, proteinaceous filaments on the bacterial surface termed type 1 pili9,10. Type 1 pili are composed of up to 3,000 copies of the subunit FimA building the pilus rod, as well as the subunits FimF, FimG and FimH forming the distal tip fibrillum11. The adhesin FimH at the fimbrial tip specifically binds in a catch-bond mode12 to terminal α-D-linked mannoses of N-linked glycans of the receptor uroplakin 1a on urinary epithelial cells13. Owing to its important role in establishing infection, FimH is an attractive target for the development of anti-adhesive drugs for UTI treatment14,15.
FimH is a two-domain protein, composed of an N-terminal, mannoside-binding lectin domain (FimHL) and a C-terminal pilin domain (FimHP). FimHP possesses an incomplete immunoglobulin-like fold that is completed by insertion of an N-terminal donor strand of FimG, the subsequent subunit in pilus assembly11. The two-domain architecture of FimH is a prerequisite for catch-bond formation because the interactions between FimHL and FimHP determine the conformational state and ligand-binding properties of FimHL (refs 12, 16, 17). A ‘compressed’ FimHL conformation was observed in the crystal structure of FimH in the context of the type 1 pilus tip fibrillum in the absence of ligands, with an open binding site and interactions to FimHP mediated via three loop segments: the swing (amino acids (aa.) 27–33), linker (aa. 154–160) and insertion loops (aa. 112–118)17. In contrast, an ‘extended’ FimHL conformation was observed in crystal structures of the isolated, ligand-bound FimHL domain18,19,20,21,22,23 and in the complex between FimH and the pilus assembly chaperone FimC, where FimC prevents the interactions between FimHL and FimHP (ref. 24). This extended form of FimHL is characterized by a closed ligand-binding pocket and rearranged swing, linker and insertion loops.
Notably, isolated FimHL was reported to show a ligand-binding affinity about two orders of magnitude higher than that of full-length FimH in the tip fibrillum17,25. Together with mutagenesis experiments disrupting the interdomain interface26, these data indicated that ligand-binding is linked to domain separation in FimH, and that mechanical force shifts the ligand-binding affinity towards that of the isolated FimHL. However, fundamental aspects of the mechanism underlying the force-dependent binding of FimH remained unknown: (i) How is domain-associated, full-length FimH interacting with ligands? (ii) Does ligand-binding directly induce domain separation? (iii) How are interdomain interactions linked to the ligand-binding affinity of FimH and the kinetics of ligand-binding and dissociation?
To address these questions, we designed a stable, soluble variant of full-length FimH that is equivalent in its structural and functional properties to those of FimH in the assembled fimbrial tip. This variant allowed us to obtain high-resolution structural snapshots of all functional states of FimH and to obtain a complete characterization of ligand-binding kinetics in solution. Together with molecular dynamics simulations, these data reveal a three-state mechanism of FimH catch-bond formation. FimHP accelerates ligand release from FimHL via dynamic allostery by 100,000-fold. In addition, using single-cell tracking experiments, we show that the modulation of ligand affinity by FimHP is not only required for adhesion under mechanical stress, but also for efficient bacterial surface motility in the absence of shear force. Our results provide a first complete structural and kinetic description of a catch-bond system and establish a framework for the analysis of the distinct catch-bond mechanisms in other systems, which also commonly couple interdomain interactions to ligand affinity.
Construction of a peptide-complemented FimH
Isolated FimH with its non-complemented pilin domain is only marginally stable and shows aggregation tendency under physiological conditions27. To establish a stable, isolated FimH molecule with all properties of FimH in the tip fibrillum, we complemented FimHP with the donor-strand peptide of FimG (FimG residues 1−14; termed DsG). The FimH·DsG complex was obtained in good yields and purified after an in vitro reaction, mimicking the first donor-strand exchange (DSE) reaction during pilus assembly in vivo. In this reaction, the FimG donor strand displaces the pilus assembly chaperone FimC from FimH (Fig. 1a):
The experiments described in the following were performed with FimH from the faecal E. coli strain F18 (FimHF18), which is structurally identical to the most prevalent variants in uropathogenic infection25, and FimH from the wild-type E. coli strain K12 (FimHK12), which differ in three amino acids in FimHL (K12→F18: Val27Ala, Asn70Ser, Ser78Asn; Supplementary Fig. 1a). The isolated lectin domains (residues 1–159) of both FimH variants (FimHLK12 and FimHLF18) were produced by direct expression in the E. coli periplasm and were purified as described27.
Ligand-free FimH·DsG resembles FimH in the fimbrial tip
The crystal structure of the binary complex FimHF18·DsG was determined at atomic resolution by molecular replacement (Table 1). FimHF18·DsG comprises the jellyroll fold FimHL and the immunoglobulin-like FimHP domain complemented with the FimG donor strand (Fig. 1b and Supplementary Fig. 1b). It closely resembles unliganded FimH in the fimbrial tip complex (Fig. 1c)17, with a root-mean-square deviation of Cα positions (Cα r.m.s.d.) of 1.1 Å. The individual FimHP and FimHL domains are even more closely resembling unliganded, fimbrial FimH (r.m.s.d. 0.45 and 0.55 Å, respectively) and undergo only a minimal hinge-bending rotation of 4° (Fig. 1c).
The DsG peptide in FimHF18·DsG is in identical position as compared with the N-terminal FimG extension in the fimbrial tip structure; it interacts with β-strand 2 and 9 of FimHP (Fig. 1d). All contacts in the FimHL–FimHP interdomain region (Supplementary Fig. 1c,d)17 as well as the conformation of the empty ligand-binding pocket observed in FimH in the fimbrial tip are preserved in FimHF18·DsG. Thus, FimHF18·DsG represents the ligand-free state of fimbrial FimH, with Associated FimHL and FimHP (Afree state) and is an elegant minimal system to analyse the crosstalk between ligand-binding and interdomain interactions underlying the formation of catch-bonds by FimH.
Persistence of domain association in ligand-bound FimH·DsG
To test whether ligand-binding causes domain separation in FimH, we determined the co-crystal structure of the ternary complex of FimHF18·DsG with n-heptyl α-D-mannoside (HM), an established model ligand of FimH20, as well as crystal structures of the isolated FimHLF18 and FimHLK12 lectin domains in complex with HM (Table 1). FimHF18·DsG·HM adopts the same closed conformation of the ligand-binding site as previously observed in other FimHL–ligand complexes (Fig. 2a,b and Supplementary Fig. 2)28. The mannopyranose moiety of HM is coordinated by the side chains of Asp54, Gln133, Asn135 and Asp140, and the main chain of Phe1 and Asp47, and the n-heptyl aglycone of HM is sandwiched between Tyr48 and Tyr137. Compared with the Afree form, all loops surrounding the binding pocket close down onto the HM ligand. The most substantial conformational difference to Afree is observed for the clamp loop (aa. 8–16), whose tip moves almost 6 Å towards HM (Supplementary Fig. 1e,f).
Besides the closing of the ligand-binding pocket, the overall conformation of ligand-free FimHL in Afree and HM-bound FimHF18·DsG is closely similar (Cα r.m.s.d. 1.1 Å; Fig. 2a,b and Supplementary Fig. 2). Unexpectedly, the structural change in the ligand-binding site in FimHF18·DsG·HM was not transmitted to the domain interface, where the interdomain contacts and the conformations of the swing, linker and insertion loops remained intact. The lectin domain in the FimHF18·DsG·HM complex thus differs drastically from HM-bound isolated FimHL domains with respect to the swing, linker and insertion loop conformations (Fig. 2c). On the basis of the persistence of the domain Association in the ligand-bound form, this state of FimH was termed Abound.
To test the stability of the Abound state against domain separation and to exclude potential effects of selective crystallization, a molecular dynamics (MD) simulation of the Abound state was conducted using the CHARMM36 force field of the NAMD package (Supplementary Fig. 3a,b). The domain association remained intact over 100 ns of simulation time without substantial changes in the domain interface; fluctuations were limited to the clamp loop region close to the ligand-binding site. On in silico removal of the HM ligand after initial equilibration, the Abound state underwent a spontaneous transition to the Afree state after ∼75 ns of simulation time via an opening of the clamp loop (Supplementary Fig. 3c,d), reproducing the experimentally observed dependence of the binding-site conformation on ligand-binding. Thus, the MD simulations indicate that Abound is a stable conformational state of FimH induced by ligand-binding.
Trapping of a domain-separated state of full-length FimH
The increase in apparent affinity of FimH to its target glycans under tensile mechanical forces12,29 has previously been linked to a separation of the FimHL and FimHP (ref. 17). To trap a potential domain-separated state of FimH for structural characterization in the absence of tensile force, we considered FimH variants with weakened interdomain interactions. We had shown previously that FimHP also accepts the donor strand of the non-cognate subunit FimF (DsF). However, FimHP is slightly less stabilized by complementation with DsF than with the natural donor-strand DsG30. We hypothesized that such complementation with DsF instead of DsG could also result in a mild destabilization of the interdomain interface in full-length FimH.
We then determined the co-crystal structure of FimHK12·DsF with HM (FimHK12·DsF·HM) at 3.0 Å resolution with four molecules in the asymmetric unit. Three FimHK12·DsF·HM molecules closely resembled the Abound state (r.m.s.d. of 0.6 Å to FimHF18·DsG·HM) with a preserved interdomain interface. In the fourth molecule, however, the FimHL and FimHP domains were separated and they adopted a drastically different relative orientation with an angle between the domains of ∼45° instead of ∼150° in the other three molecules (Fig. 2a). FimHP is virtually identical in all four FimH molecules in the crystal (r.m.s.d. 0.4 Å). In contrast, the FimHL domain differs significantly between the fourth, domain-separated and the three full-length FimH molecules in the crystal. It shows closest similarity to the isolated FimHL·HM (r.m.s.d. 0.45 Å); in particular, all interdomain loops adopt identical conformations, which are incompatible with domain association (Figs 2c and 3). Remarkably, in the bent fourth molecule, no interactions between FimHL and FimHP other than the direct covalent linkage are detected, equivalent to a breakdown of the total 500 Å2 interdomain interface of the Abound state (Fig. 3). This molecule thus represents a third state, the domain-Separated, ligand-bound state of FimH, Sbound. The complete absence of non-covalent interdomain interactions indicates that the Sbound state does not possess a defined relative domain orientation in solution, and that the observed, kinked conformation has been selected only by crystal packing.
To analyse the transition trajectory of the Abound to the Sbound state, we removed the FimHP domain after equilibration from the Abound state in silico for a 180-ns molecular dynamics simulation (Supplementary Fig. 3e,f). In contrast to the transition between the Abound and Afree states on ligand removal, a sharp transition to the conformation of FimHL in the Sbound state was not observed. The conformation only slowly moved towards Sbound; however, the FimHL loops that had formed in the former interdomain interaction kept fluctuating throughout the simulation, indicating lower cooperativity and potentially a higher activation energy for the Abound→Sbound compared with the Abound→Afree transition.
A comparison of the structural dynamics in the Abound and Sbound states clearly reveals differences in the FimHL–FimHP interface region. The root-mean-square fluctuations of atom positions (r.m.s.f.) increase in the swing and insertion loop from a background level of ∼0.7 Å in Abound to 1.5 and 2 Å in Sbound, respectively. Surprisingly, despite the virtually identical conformations of the entire ligand-binding site depicted by X-ray crystallography (Fig. 2b), the clamp loop, which exhibits the most significant conformational changes between the open and closed conformations, exhibits strongly reduced fluctuations in Sbound, with r.m.s.f. decreasing by up to 1.5 Å (Supplementary Fig. 3g,h). This change in clamp loop dynamics provides a mechanistic link between domain association and ligand-binding in full-length FimH.
Domain association alters FimH–ligand-binding kinetics
To analyse the ligand-binding properties of FimH·DsG, we exploited the increase in intrinsic tryptophan fluorescence in the FimH·DsG complexes of ∼10% on HM binding (Fig. 4a). This difference was used to measure the dissociation constant of HM binding by equilibrium titration (Fig. 4b) and the rates of HM binding and dissociation by stopped-flow fluorescence kinetics (Fig. 4c,d). The FimH·DsG constructs showed uniform binding and dissociation kinetics, consistent with the view that domain-separated states of FimH are not significantly populated in the absence of shear force. The results revealed equilibrium dissociation constants (Kd) of 3.6 and 9.9 μM for FimHK12·DsG and FimHF18·DsG, respectively (Table 2). HM binding to FimH·DsG is extremely dynamic and was characterized by fast association rates (kon) of 5.0 × 106 and 4.9 × 106M−1s−1, respectively, and rapid dissociation reactions (Supplementary Fig. 4). The rates of HM dissociation (koff) of 22 and 58 s−1 for FimHK12·DsG and FimHF18·DsG translate into dissociation half-lives of only 32 and 12 ms, respectively.
In contrast to full-length FimH, isolated FimHLK12 showed no change in tryptophan fluorescence on HM binding. We therefore determined the HM affinity of isolated FimHL indirectly by a competition experiment based on a newly designed fluorescent ligand, the fluorescein-labelled α-D-mannoside GN-FP-4 (Supplementary Fig. 5a–e and Supplementary Note 1). Displacement of GN-FP-4 from FimHL by increasing HM concentrations under equilibrium conditions showed that both FimHLK12 and FimHLF18 bind HM with 3,300-fold higher affinity compared with the respective FimH·DsG complexes (Kd values 1.1 and 3.0 nM, respectively; Fig. 5a and Table 2). In an inverse competition experiment (Supplementary Fig. 5f–j), in which HM in preformed FimHL·HM complexes was displaced by GN-FP-4, off-rates of 2.0 × 10−4 and 3.5 × 10−4s−1 were determined for FimHLK12 and FimHLF18, respectively, corresponding to dissociation half-lives of 58 and 33 min (Table 2). On the basis of these measured off-rates and equilibrium dissociation constants, kon rates of 1.8 × 105 and 1.2 × 105M−1s−1 were calculated for FimHLK12 and FimHLF18, respectively. The on-rates for the isolated FimHL domains are thus 30-fold lower than those of the corresponding full-length FimH·DsG complexes.
Together, these results demonstrate that the 3,300-fold higher affinity of the isolated FimHL compared with full-length FimH results from a more than 100,000-fold lower ligand dissociation rate in isolated FimHL, combined with a ligand-binding rate reduced by only 30-fold (Table 3). The 3,300-fold higher affinity for HM of FimHL relative to FimH·DsG translates into a free energy of 20 kJ mol−1 for the interaction between FimHL and FimHP in full-length FimH. This corresponds very well with the mechanical work required for domain separation, as a displacement of FimHL from FimHP by 11 Å for complete domain separation (Fig. 2a)17, and a force of 40 pN required to populate the domain-separated state of FimH31 yields a value of 26.5 kJ mol−1.
Domain association in FimH promotes bacterial motility
Uropathogenic E. coli require firm adhesion to the urinary epithelium under flow conditions to escape clearance by urine excretion. On the other hand, bacterial adhesion must be weak enough in the absence of external shear to allow flagellar motility as a prerequisite for the invasion of new tissue areas32,33. While the role of FimH catch-bond binding for adhesion under flow conditions had clearly been demonstrated12,29,34, the relevance of rapid ligand dissociation under static conditions for flagellar motility remained unclear because of the complex interplay of flagellar swimming and the avidity of multivalent surface interactions by hundreds of E. coli pili. Here we employed single-cell tracking of piliated E. coli cells moving on surfaces coated with mono-mannosylated bovine serum albumin (1M-BSA), an established model system for analysing FimH-based adhesion12,29, for a classification of cell motility into two states, attached or mobile (for details see Methods and Supplementary Fig. 6). To study the influence of FimH interdomain interactions, we compared isogenic E. coli strains producing either wild-type FimHF18 or the FimHF18-variant Ala188Asp, which is characterized by a destabilized interaction between FimHP and FimHL (ref. 26) and serves here to mimic the Sbound state in the absence of shear force35. The overall fraction of adherent FimHF18-piliated bacteria on 1M-BSA-coated surfaces was identical to background levels on non-adhesive BSA-coated surfaces at 10–12% of tracked bacteria (Fig. 6a). In contrast, FimHF18–Ala188Asp-piliated bacteria showed an increased fraction of adherent cells of 24% already at the beginning of cell tracking after a 1-min dead time, which further increased during the 5-min observation period to 48% (Fig. 6a and Supplementary Movies 1 and 2).
Cell tracking permitted quantitative analysis of the transition of individual cells between a mobile and an attached state (Supplementary Fig. 7). On non-adhesive control surfaces coated only with BSA, less than 2% of all FimHF18- or FimHF18–Ala188Asp bacteria showed shifts between the two states of motion (Fig. 6b and Supplementary Fig. 7c). However, on adhesive 1M-BSA surfaces, 13.9% of all FimHF18 tracks (green in Fig. 6b) exhibited a single transient attachment event with a mean duration of 6.9 s (Supplementary Fig. 7d). For FimHF18–Ala188Asp-piliated bacteria, only 7.2% of the cells showed attachment/detachment, but with fivefold longer adhesion (35.2 s; Supplementary Fig. 7d). Remarkably, the fraction of cells that permanently stayed attached after adhesion to 1M-BSA until the end of the observation period was much larger for FimHF18–Ala188Asp (11.5%) than for FimHF18 (0.6%; red in Fig. 6b). Those permanently attached cells escape kinetic analysis; thus, the true average attachment time for FimHF18–Ala188Asp must be considerably larger than 35.2 s. Permanent attachment is also the main cause of the increased fraction of attached cells for FimHF18–Ala188Asp-piliated bacteria (Fig. 6a).
Altogether, cell-tracking analysis revealed that enforced domain separation in the FimHF18–Ala188Asp variant resulted in reduced detachment rates and a larger proportion of permanently attached cells. These results directly demonstrate, at the cellular level, the importance of fast, spontaneous ligand dissociation catalysed by interdomain allostery in FimH–ligand complexes for bacterial motility in the absence of tensile mechanical forces.
The characterization of full-length FimH had so far been restricted to the analysis of the adhesive properties of piliated E. coli cells and binding studies with the purified type 1 pilus tip fibrillum. With the FimH·DsG complex, we have now established a model system for quantitative studies of the interaction of FimH with carbohydrate ligands. Soluble FimH·DsG efficiently mimicks FimH in the context of the assembled tip fibrillum, is readily available in milligram quantities and permits the determination of ligand-binding and release kinetics in solution. Using FimH·DsG, we obtained high-resolution snapshots of three functionally relevant states of FimH (Fig. 7). In the absence of ligands, FimH adopts the Afree state with associated FimHL and FimHP and an open conformation of the ligand-binding site, which is responsible for the 30-fold faster ligand-binding of full-length FimH as compared with the isolated FimHL domain. Ligand-binding in the absence of shear force induces the Abound state with a closed binding site. In contrast to earlier hypotheses17, the transition from Afree to Abound is restricted to the ligand-binding site, while all interactions between FimHL and FimHP observed in the Afree state remain preserved in Abound. The Afree→Abound transition most likely follows an induced fit mechanism, in which the formation of an encounter complex between FimH·DsG and HM is rate-limiting and followed by a fast, unimolecular rearrangement to the Abound state, in agreement with the observation that binding of the model ligand HM remained rate-limiting for the formation of Abound even at the highest HM concentrations used. Stopped-flow-binding kinetics indicate that the lifetime of the proposed encounter complex before Abound formation is below 1 ms (Fig. 4c). Under tensile mechanical force applied to the FimH–ligand complex, mimicked here by the destabilized variant FimH·DsF and crystal packing forces, the domain-separated state of FimH, Sbound, is formed. In the Sbound state, FimHL and FimHP no longer interact specifically and are only connected via the linker segment comprising FimH residues 154–160. In this Sbound state, FimHL adopts a conformation closely resembling isolated FimHL with bound ligand.
Notably, ligand dissociation from FimH·DsG is 100,000-fold faster than that from the isolated FimHL domain. This is striking because the respective crystal structures revealed indistinguishable ligand interactions and binding-site conformations in the FimH·DsG·HM and FimHL·HM complexes (Fig. 2b). MD simulations identified a considerable increase in the conformational dynamics of the FimHL–ligand-enclosing clamp loop in the Abound state as the most likely cause of the dramatic increase in koff in the FimH·DsG·HM complex. The altered dynamics in FimHL in the Abound state are the result of the presence of FimHP, which can be described as a negative allosteric regulator36,37,38. The allosteric communication from the FimHP–FimHL interface to the ligand-binding site reaches over 40 Å, and is mediated via changes in protein dynamics rather than in static structure, in line with a general model of dynamic allostery39,40. Our data demonstrate that the interdomain interactions in FimH (i) maintain the open conformation of the binding pocket and guarantee rapid ligand-binding and (ii) intramolecularly catalyse ligand dissociation by more than 100,000-fold. Rapid ligand-binding and short lifetimes of the FimH ligand complex allow for rapid dissociation of individual pili from their ligands in the absence of shear force. Our biophysical data demonstrate that this mechanism is conserved between the K12 and the F18 E. coli strains.
Different mechanistic models, such as the two-pathway41, the deformation42 and the sliding re-binding model43, have been developed to describe catch-bond interactions, often based on powerful single-molecule atomic force measurements. These models included the principle of allosteric control of ligand-binding affinity26,31, which was clearly fully confirmed in our present study. However, these conceptual models did not reveal the underlying atomic-scale mechanisms in different catch-bond systems. For most catch-bond systems, including the cadherin–catenin binding to actin filaments3,44, integrin epithelial cell adhesion45,46 and TCR–MHC interactions6,7,47, structural information is, if at all, available only for one state or from computer simulations. One exception is the selectins, which employ catch-bond binding for leukocyte recruitment. Selectins are multidomain cell surface receptors, which consist of a lectin domain for complex carbohydrate binding, linked via an epidermal growth factor (EGF)-like domain to a variable number of short consensus repeat domains and a transmembrane-anchoring helix. Selectins exist in two conformations, a bent and an extended one, which differ in the angle between their lectin and EGF-like domain. Ligand-binding and conformational changes in the ligand-binding site are directly linked via a complex allosteric coupling mechanism to the adoption of the extended conformation48,49. Tensile mechanical force under flow conditions acts along the axis of the ligand-binding site and the Lec-EGF interface resulting in a stabilization of the extended conformation and thus increased ligand complex lifetimes2,49. Moreover, in FimH, catch-bond behaviour is mediated by the interplay of a lectin and an anchoring domain that does not interact with the ligand. Ligand-binding by FimH in the absence of shear force results in a closing of the ligand-binding site, but, in contrast to selectins, is not directly linked to altered interdomain interactions. Here mechanical force promotes domain separation and completely releases FimHL from FimHP, which acts as an activator of ligand release via dynamic allostery. Remarkably, selectins and the fimbrial adhesin FimH thus employ entirely different mechanisms for establishing catch-bond behaviour by crosstalk between a lectin and an anchoring domain that provides tethering to a shaft. In both systems, the selectins and fimbrial adhesion, the shaft structures linking the terminal lectin/coupling domains to the cell surface, may contribute to the overall catch-bond behaviour, either via directly influencing coupling domain behaviour or via their general elastic properties50,51.
The cell-tracking experiments indicate the importance of rapid ligand release from the high-mannose-type glycoprotein receptor uroplakin 1a in the lower urinary tract52 for flagellar motility of piliated bacteria, and hence their ability to colonize new tissue areas under certain conditions during infection12,29,53. This provides a plausible explanation for the fact that low-affinity FimH variants were preserved in numerous uropathogenic E. coli strains. Binding of terminal mannoses with low affinity in the absence of shear force may also play a role in preventing the clearance of uropathogenic E. coli from the urinary tract by competitive binding to the Tamm–Horsfall protein in the urine54. In turn, populating the Sbound state with an extremely low dissociation rate ensures tight bacterial adhesion under the mechanical forces of urine excretion. FimH is a promising target for anti-adhesive therapy of UTI because FimH antagonists, in contrast to antibiotics, are not exerting selection pressure towards resistance formation18,55,56. Previous ligand-binding studies on the isolated FimHL domain mimic the domain-separated Sbound state of FimH. This state is characterized by extremely low off-rates and is promoted in vivo only after ligand-binding and the onset of flow conditions. Our kinetic data on ligand dissociation from full-length FimH demonstrate that rapid, competitive displacement of FimH from its carbohydrate ligands by FimH antagonists is well possible in the absence of shear force. Thus, full-length FimH (for example, in the form of the FimH·DsG complex established in this study) instead of the isolated FimHL domain is the relevant target for the development of anti-adhesive drugs. Importantly, the concept of the FimH·DsG model system can now be expanded to other related adhesive pilus adhesins. In combination with the novel fluorescent GN-FP-4 ligand, this model system paves the way for efficient screening for anti-adhesive drug candidates.
The synthetic DsG (sequence: ADVTITVNGKVVAKR) and DsF peptide (sequence: ADSTITIRGYVRDNG; >95% purity) were purchased from JPT (Germany). Guanidinium chloride (‘AA-Grade’ for spectroscopy) was obtained from NIGU Chemie (Germany). Standard chemical of highest purity available was obtained Sigma, Merck or AppliChem. If not mentioned otherwise, chromatography media for protein purification were purchased from GE Healthcare (UK). Oligonucleotides were from Microsynth (Switzerland).
Construction of expression plasmids
Expression plasmids for the periplasmic production of the E. coli F18 FimH lectin domain (FimHLF18) and for the periplasmic co-expression of full-length FimHF18 with FimC were based on the expression plasmids pfimHL and pfimH-fimC-ATG, respectively, for the analogous proteins from E. coli K12 (ref. 27). Six silent mutations replacing rare codons were introduced into the E. coli F18 fimH gene (fimHF18) contained in the plasmid pGB2-24 (ref. 57) with the QuikChange mutagenesis kit (Agilent Technologies, Switzerland) to improve periplasmic expression. The coding sequence of the modified fimHF18 gene was amplified by PCR using the primers 5′-GATCCTCTAGAGGAGGGATGATTGTAATGAAACGAG-3′ and 5′-TTTCAAGCTTATTGATAAACAAAAGTCACG-3′ and cloned into pfimH-fimC-ATG27 via the XbaI and HindIII sites (thereby replacing the fimHK12 gene) and yielded pfimHF18–fimC–ATG. The gene encoding FimHLF18 was amplified with the primers 5′-GATCCTCTAGAGGAGGGATGATTGTAATGAAACGAG-3′ and 5′-CAGCCAAGCTTAGCCAGTAGGCACCACCAC-3′ and ligated via the XbaI and HindIII sites into ptrc99a-f1-stopp27. Protein production in the resulting plasmid pfimHLF18 is under control of the trc–promotor/lac operator.
Protein production and purification
For purification of the complexes FimC·FimHK12 and FimC·FimHF18, E. coli HM125 harbouring the corresponding co-expression plasmid was grown at 30 °C in 2YT medium containing ampicillin (100 μg ml−1). At an OD600 of 1.5, isopropyl-β-D-thiogalactoside (IPTG) was added to a final concentration of 1 mM. The cells were further grown for 12−18 h, harvested by centrifugation, suspended in cold 50 mM Tris-HCl pH 7.5, 150 mM NaCl, 5 mM EDTA, 1 mg ml−1 polymyxin B sulfate (18 ml l−1 of culture) and stirred at 4 °C for 1.5 h. After centrifugation, the supernatant (periplasmic extract) was dialysed against 20 mM Tris-HCl pH 8.0 and applied to a QA52 (Whatman, Maidstone, UK) column equilibrated with the same buffer. The flow-through containing the respective FimC·FimH complex was dialysed against 20 mM MOPS–NaOH pH 7.0, loaded onto a Resource S column equilibrated with the same buffer and the complexes were eluted with a linear NaCl gradient (0−400 mM). Fractions containing FimC·FimH were pooled and loaded onto a Superdex 75 (HiLoad 26/60) column equilibrated with 20 mM NaH2PO4–NaOH pH 7.4, 50 mM NaCl. Fractions containing the pure complex were pooled and stored at 4 °C until further use. Typically, 3−5 mg of the purified complex were obtained per litre of bacterial culture.
For expression of the isolated E. coli FimHLK12 and FimHLF18, E. coli HM125 transformed with the respective expression plasmid was grown at 30 °C in M9 medium containing ampicillin (100 μg ml−1) to an OD600 of 1.0, and expression was induced with 1 μM IPTG. After further growth for 12 h, cells were subjected to periplasmic extraction (see above). The extracts were mixed with 0.11 volumes of 1 M acetic acid–NaOH pH 4.5, dialysed against 10 mM acetic acid–NaOH pH 4.5 and then loaded onto a SP-Sepharose column equilibrated with the same buffer. The flow-through was collected and its pH was adjusted to 8.0 by addition of 1 M Tris-HCl pH 8.2. This solution was then applied to a Q-Sepharose column equilibrated with 20 mM Tris-HCl pH 8.0. The flow-through containing FimHL was loaded onto a Resource S column dialysed against 20 mM formic acid–NaOH pH 4.0. The protein was eluted with a linear NaCl gradient (0−1 M). Fractions containing pure FimHL were pooled, dialysed against water and stored at −20 °C. The identity of the purified proteins was confirmed by electrospray ionization (ESI)–mass spectrometry (FimHLF18: calculated: 16,934.9 Da; measured: 16,935.0 Da; FimHLK12: calculated: 16,963.0 Da; measured: 16,962.8 Da). About 11 mg of the pure FimHL was obtained per litre of bacterial culture.
Production of FimH·DsG and FimHK12·DsF complexes
The respective FimC·FimH complex (40 μM) was incubated with a threefold molar excess of the DsG peptide and incubated in 20 mM NaH2PO4–NaOH, pH 7.0, 50 mM NaCl for 48 h at 37 °C. The reaction mixture containing isolated FimC, the FimH·DsG complex and excess DsG was dialysed against 20 mM acetic acid–NaOH pH 4.5 and loaded onto a Resource S (6 ml) column equilibrated with the same buffer. The FimH·DsG complex was eluted with a linear NaCl gradient (0−400 mM). Fractions containing the pure complex were pooled, dialysed against water and stored at 4 °C. The FimH·DsG partially dissociated during ESI–mass spectrometry analysis, so that masses of the intact complexes and free FimH were obtained: FimHK12·DsG: calculated mass: 30,635.3 Da; measured mass: 30,636.0 Da; FimHF18·DsG: calculated mass: 30,607.3 Da; measured mass: 30,607.0 Da; FimHK12: calculated mass: 29,064.5 Da; measured mass: 29,064.0 Da; FimHF18: calculated mass: 29,036.4 Da; measured mass: 29,036.0 Da. The overall yields of the purified FimH·DsG complexes relative to the initial amount of FimC·FimH were in the range of 50–55%. The FimHK12·DsF complex was generated and purified as described for the FimH·DsG complexes (FimHK12·DsF: calculated mass: 30,702.2 Da; measured mass: 30,702.5 Da). The FimH·DsF complex was prepared from the FimC·FimH complex after mixing with DsF exactly according to the protocol described above for FimH·DsG and obtained in similar yields. Despite the non-natural interaction between FimHP and DsF, the FimHK12·DsF complex was formed four times faster than the FimHK12·DsG complex at pH 7.0 and 37 °C, with a rate constant of 2.2±0.5 M−1 s−1. The FimH·DsF complex was stable against dissociation and unspecific aggregation.
Determination of protein concentrations
Protein concentrations were measured via the specific absorbance at 280 nm, using the following extinction coefficients (FimHK12 and FimHF18 have identical extinction coefficients): FimC·FimH (59,090 M−1 cm−1), FimHL (24,670 M−1 cm−1), FimH·DsG (35,090 M−1 cm−1) and FimH·DsF (36,580 M−1 cm−1). The concentrations of DsG and DsF were determined via their absorbance at 205 nm (42,650 and 49,700 M−1 cm−1, respectively).
Synthesis of the fluorescent-labelled FimH ligand GN-FP-4
To a stirred solution of mannoside 1 (25 mg, 0.061 mmol)55 in dry dimethylformamide (DMF; 1 ml), N-hydroxysuccinimide (21 mg, 0.183 mmol) was added, followed by N,N′-dicyclohexylcarbodiimide (9.2 mg, 0.073 mmol). The mixture was stirred at room temperature for 2 h, then N-Boc-ethylendiamine (10.7 mg, 0.067 mmol) was added and the reaction was stirred for an additional 10 h. After cooling to 0 °C, the reaction mixture was diluted with water and concentrated. Chromatography on silica gel (CH2Cl2/MeOH) yielded 23 mg (0.042 mmol, 68%) of tert-butyl (3′-chloro-4′-(α-D-mannopyranosyloxy)-biphenyl-4-yl-carboxamido)ethyl)carbamate. This product was dissolved in CH2Cl2 (3 ml) and trifluoroacetic acid (TFA, 1 ml) was added. The solid dissolved during addition of TFA. After 10 min at room temperature the reaction was complete. The mixture was evaporated and excess TFA was removed in high vacuum. The intermediate N-(2-aminoethyl)-3′-chloro-4′-(α-D-mannopyranosyloxy)-biphenyl-4-carboxamide TFA salt (23 mg, 0.042 mmol, quant.) was used without purification in the next step. After dissolution in dry DMF (0.5 ml), triethylamine (12.8 mg, 0.127 mmol) was added. The mixture was cooled to 0 °C, then fluorescein isocyanate (14.8 mg, 0.038 mmol) was added and the mixture was stirred for 3 h in the dark. After the addition of water, DMF was removed azeotropically, the residue dissolved in MeOH/10% acetic acid and evaporated. Chromatography on silica gel (CH2Cl2/MeOH) yielded compound 2, contaminated with triethylammonium acetate. Therefore, after dissolution in MeOH, 0.5 N HCl in MeOH was added, the mixture evaporated and chromatographed on silica gel to yield pure compound GN-FP-4 ((3′-Chloro-N-(2-(3-(3′,6′-dihydroxy-3-oxo-3H-spiro[isobenzofuran-1,9′-xanthen]-5-yl)-thioureido)ethyl)-4′-(α-D-mannopyranosyloxy)-biphenyl-4-carboxamide)58 (15 mg, 47%). [α]D20+12.1 (c 0.3, MeOH); 1H NMR (500 MHz, CD3OD): δ=8.12 (s, 1H), 7.92 (d, J=8.3 Hz, 2H, Ar–H), 7.70 (dd, J=5.0, 13.1 Hz, 2H, Ar–H), 7.64 (d, J=8.3 Hz, 2H, Ar–H), 7.54 (dd, J=2.2, 8.6 Hz, 1H, Ar–H), 7.46 (d, J=8.7 Hz, 1H, Ar–H), 7.09 (d, J=8.2 Hz, 1H, Ar–H), 6.74 (s, 2H), 6.69 (d, J=1.4 Hz, 2H, Ar–H), 6.55 (d, J=8.4 Hz, 2H, Ar–H), 5.63 (d, J=1.3 Hz, H-1), 4.15 (dd, J=1.8, 3.1 Hz, H-2), 4.03 (dd, J=3.4, 9.5 Hz, H-3), 3.94 (s, 2H, CH2), 3.86-3.64 (m, 6H, H-4, H-5, H-6, CH2); 13C NMR (126 MHz, CD3OD): δ=153.21, 143.84, 136.41, 129.66, 129.18, 127.76, 127.70, 125.37, 118.64, 103.62 (Ar-C), 100.75 (C-1), 76.00 (C-5), 72.41 (C-3), 71.86 (C-2), 68.24 (C-4), 62.69 (C-6), 40.76 (CH2); ESI–MS: m/z: Calculated for C42H37ClN3O12S [M+H]+: 842.2, found: 842.2.
Fluorescence emission spectra of FimH variants were recorded between 300 and 450 nm (excitation at 280 nm) at 25 °C in 1.0 × 0.4-cm quartz cuvettes on a QM 7/2003 spectrofluorimeter (PTI, USA) equipped with a magnetic stirrer. Protein concentrations were 1−2 μM in 20 mM MOPS–NaOH pH 7.4. Fluorescence spectra of GN-FP-4 (ɛ495 nm=54,900 M−1 cm−1) were recorded between 500 and 650 nm (excitation at 497 nm) in the same buffer.
Kinetics of HM binding to FimH·DsG
The rate constants of binding (kon) and dissociation (koff) for the complex between FimH·DsG and HM were measured at 25 °C in 20 mM MOPS–NaOH pH 7.4 in a SX20 stopped-flow instrument (Applied Photophysics, UK). A constant FimH·DsG concentration of 1 or 2 μM was used. FimH·DsG was mixed with different concentrations of HM (2−100 μM), and binding was monitored by the increase in fluorescence above 320 nm (excitation at 280 nm). The fluorescence traces were globally fitted with Dynafit59 according to a second-order binding and first-order dissociation reaction. As an additional control, the fluorescence amplitudes of the individual reactions were plotted against the total HM concentration and fitted according to equation (2). The deduced dissociation constants reproduced the Kd values obtained with equilibrium titration within experimental error.
Equilibrium titration of FimH·DsG with HM
The binding equilibrium between FimH·DsG and HM was followed at 25 °C in 20 mM MOPS–NaOH pH 7.4 on a QM 7/2003 spectrofluorometer (PTI) by the increase in fluorescence at 320 nm on HM binding (excitation at 280 nm). Measurements were performed with a stirred 1 × 0.4-cm quartz cuvette. The concentration of FimH·DsG was kept constant at 2 μM and the concentration of HM was varied between 0 and 200 μM. The samples were equilibrated overnight, and their fluorescence intensities were recorded for 30 s and averaged. The fluorescence intensities were plotted against total HM concentration and fitted according to equation (2)
where F is the monitored fluorescence signal, F0 is the fluorescence signal in absence of ligand, F∞ is the fluorescence signal at full saturation with ligand, Kd is the dissociation constant, [P]0 is the total concentration of FimH·DsG and [L]0 is the total concentration of HM.
Equilibrium titration of FimHL with GN-FP-4
The binding equilibrium between FimHL and GN-FP-4 at 25 °C in 20 mM MOPS–NaOH pH 7.4, supplemented with 0.001% Tween 20 to prevent unspecific adsorption effects at nanomolar concentrations, was recorded by the decrease in GN-FP-4 fluorescence at 520 nm (excitation at 497 nm). Measurements were performed with a stirred 1 × 0.4-cm quartz cuvette. The concentration of GN-FP-4 was kept constant at 1.0 or 2.0 nM and the concentration of FimHL was varied between 0 and 10 nM. The samples were equilibrated overnight, and their fluorescence intensities at 520 nm were recorded for 30 s and averaged. The experimental data were fitted according to equation (2).
Displacement of HM from the FimHL by GN-FP-4
The rate constant of dissociation (koff) for HM from FimHL at 25 °C in 20 mM MOPS–NaOH pH 7.4 was measured indirectly by binding of excess GN-FP-4 to FimHL after dissociation of HM, recorded with the decrease in GN-FP-4 fluorescence at 520 nm (excitation at 497 nm). A mixture of FimHL and HM (3 μM each), pre-incubated for at least 18 h, was mixed with different amounts of excess GN-FP-4 (final concentrations: 10−40 μM), and GN-FP-4 fluorescence was recorded every 10 min for 10 s, averaged and the data fitted according to first-order kinetics. The obtained rate constants were independent of GN-FP-4 concentration and thus identical to the dissociation rate of HM from FimHL.
Determination of the FimHL·HM dissociation constant
The affinity of FimHL for HM at 25 °C in 20 mM MOPS–NaOH pH 7.4 was determined by the competition between HM and GN-FP-4 for binding to FimHL. A mixture of FimHL and GN-FP-4 (1 μM each) was incubated with different concentrations of HM (10–3.2 mM) and incubated for at least 18 h. The displacement of GN-FP-4 by HM was recorded on by the decrease in the fluorescence polarization at 528±20 nm (excitation at 485±20 nm) on a microplate reader (Biotek, USA), using flat black-bottom 96-well microtitre plates (Greiner, Austria). The fluorescence polarization data were fitted with Dynafit59 according to an equilibrium competition mechanism, with the total concentrations of FimHL, GN-FP-4 and HM (variable) and the respective Kd of GN-FP-4 (Table 2) as input, and Kd of HM and the fluorescence polarization at zero and infinite HM concentration as open parameters.
Crystallization of FimH variants
All crystallization experiments were performed at 4 °C with the sitting drop vapour diffusion method. For crystallization, FimHF18·DsG and FimHK12·DsG (0.1−0.2 μl, 15 mg ml−1 in H2O) was mixed with 0.1−0.2 μl of precipitant (25% (w/v) polyethylene glycol (PEG) 3350, 0.2 M magnesium chloride, 0.1 M BisTris-HCl pH 5.5 at 4 °C. Crystals of FimHF18·DsG and FimHK12·DsG grew within 4−6 weeks and are of the space group C2, with one molecule per asymmetric unit. FimHK12·DsG crystals in space group P1 grew at 0.2 M Na Malonate, 20% PEG3350 within 2 months at 4 °C. For crystallization of the FimHF18·DsG·HM complex, a threefold excess over FimH·DsG was used (protein concentration and protein/precipitant ratios were as described for FimH·DsG). Crystals of the space group P213 appeared after 1 month in 30% (v/v) 2-Methyl-2,4-pentanediol, 0.1 M sodium cacodylate, 0.2 M magnesium acetate pH 6.5 at 4 °C. FimHK12·DsF·HM crystals appeared after 2 months in 30% w/v PEG 5000, 0.1 M 2-(N-morpholino)ethanesulfonic acid (MES) monohydrate, 0.2 M ammonium sulfate pH 6.5 at 20 °C (2.5-fold excess of ligand over protein). FimHLF18·HM crystallized in 17% PEG 2000 MME, 0.1 M HEPES–NaOH pH 7.5 at 4 °C. FimHLK12·HM crystallized in 1.5 M (NH4)2SO4, 0.2 M Na acetate pH 5.5 at 20 °C.
Crystallographic data collection
All crystals, except for FimHF18·DsG·HM, were cryo-preserved by the addition of ethane-1,2-diol to a final concentration of 20% (v/v). The precipitant solution used for the crystallization of FimHF18·DsG·HM already contained 30% (v/v) methyl-2,4-pentanediol, which acts as cryoprotectant. Crystals were flash-cooled in liquid nitrogen. All measurements were carried out at the SLS beamline X06DA and X06SA (Swiss Light Source, Paul Scherrer Institute, Switzerland) at 100 K. All data were integrated, indexed and scaled using the XDS software package60 (5% of the reflections were set aside as test set). Data collection statistics are summarized in Table 1.
Crystallographic structure determination
All structures were solved by molecular replacement using structures of isolated FimHL (AA1-158, PDB ID: 3MCY18, and the pilin domain of FimC·FimH (AA160-297, PDB ID: 1QUN24, as search models with the programme Phaser61). Model building and structure refinement were performed with Coot (ref. 62) and PHENIX (ref. 63). Twelve out of thirteen residues could be built for the FimG donor strands in the crystal structures, and only the C-terminal lysine residue had weak electron density. Refinement statistics are summarized in Table 1.
Molecular dynamics simulations
Four molecular systems were prepared for FimHF18. The first system was constructed using the Abound state of FimHF18·DsG·HM (Supplementary Fig. 3a,b) and the second system is equivalent but HM was removed (Supplementary Fig. 3c,d). The third system contains only the FimHL and HM from the Abound X-ray structure (Supplementary Fig. 3e,f). The fourth system was prepared for FimHL and HM based on the Sbound state in the FimHK12·DsF·HM crystal structure (Supplementary Fig. 3g,h).
The CHARMM-GUI web server64 was used to prepare the molecular systems, which were solvated with TIP3 water molecules and ionized with 50 mM NaCl. Each system contains between 50,000 and 60,000 atoms. All simulations were performed with the NAMD simulation package (version 2.9) (ref. 65). The CHARMM36 force field was used for the protein, and parameters for HM were generated using the CHARMM General Force Field programme (version 0.9.7 beta). Electrostatic interactions were calculated using the particle-mesh Ewald method66 with a grid spacing of 1 Å. The cutoff for the van der Waals interactions was taken at 12 Å with a switching function used after 10 Å. Time step for the integration of dynamics was 2 fs. Simulations were performed in an isothermal–isobaric ensemble, with a pressure of 1 atm and a temperature of 300 K.
Cell tracking on mannose–BSA-coated surfaces
The E. coli KB18 strain67 was kindly provided by Professor Evgeni Sokurenko and served as host for the generation of recombinant strains. KB18 contains the pPKL114 plasmid57, which encodes the whole fim operon with a translational stop linker upstream of the fimH gene. KB18 was co-transformed with the pGB2-24 plasmid, which was isolated from the ELT115 strain and encodes fimHJ96 (kindly provided by Professor Evgeni Sokurenko). Single-nucleotide point mutations were introduced in fimHJ96 using overlap extension PCR following standard molecular techniques to obtain fimHF18 and fimHF18-Ala188Asp. The PCR products were cloned into pGB2-24 by the ApaLI and SphI sites, and KB18 was transformed with the resulting plasmid.
E. coli strains were grown from frozen stocks in LB medium supplemented with antibiotics (100 μg ml−1 ampicillin and 25 μg ml−1 chloramphenicol) until late log phase (OD600 of 1.0–1.2) and diluted to an OD600 of 0.01 before movie acquisition.
Cell culture dishes (35 mm, Corning Inc., Corning, NY) were incubated with 50 μl of 50 μg ml−1 1M-BSA in 0.02 M bicarbonate buffer for 75 min at 37 °C. The dishes were then washed three times and quenched with 0.1% PBS–BSA to remove unbound 1M-BSA and block remaining sites on the plastic surface to prevent nonspecific binding of bacteria. Controls were prepared by treating cell culture dishes with 0.1% PBS–BSA only. The bacterial suspension was added to the cell culture dishes for microscopy studies.
Cell tracking was carried out at room temperature under static conditions. A bacterial suspension of 50 μl in the late logarithmic growth phase was placed onto the cell culture dishes (diluted to OD600 of 0.01), and a cover slide was placed on top. The delay between sample placement and start of the movie acquisition was about 1 min. Time-lapse movies were recorded with a × 20 phase contrast objective using a CMOS digital camera (The Imaging Source Europe, Bremen, Germany) mounted on a Nikon Ti Eclipse inverted microscope and using the NIS Elements Basic Research software (Nikon, Zurich, Switzerland). Phase contrast images in an ∼5-μm-thick surface layer were taken at four to five frames per second over 5 min. The dead time of movie acquisition was ∼1 min. The resulting images were segmented by creating a projection of the average intensities over all frames to remove the background and by subsequent thresholding using the Maximum Entropy method in Fiji68 to obtain binary images (examples shown in Supplementary Movies 1 and 2). The segmented movies were imported into Imaris (Bitplane, Zurich, Switzerland) and tracked through the autoregressive algorithm. A time filter was applied to exclude all tracks with a length below 15 s. Tracks longer than 15 s were reviewed individually and edited manually, if necessary. Five to seven independent movies were recorded for each experimental set-up: FimHF18 or FimHF18-Ala188Asp on 1M-BSA-coated dishes and FimHF18 and FimHF18-Ala188Asp on BSA-coated cell culture dishes. E. coli piliated with FimHF18 or FimHF18-Ala188Asp binding to 1M-BSA in the absence (1,815 and 1,283 individual tracks, respectively) and in the presence of 200 μM HM (1,175 and 1,071 individual tracks, respectively) were analysed respectively. For E. coli piliated with FimHF18 or FimHF18-Ala188Asp binding to BSA 1,314 and 1,065 individual tracks, respectively, were analysed. Bacteria with a speed of <0.5 μm s−1 were classified as attached, all other bacteria were classified as mobile. Owing to limitation in the spatial and temporal resolution of movie acquisition, we did not further subdivide bacterial swimming into motility behaviours as ‘rolling’29,53, ‘roaming’, ‘orbiting’ and so on. The individual cell tracks were classified into four classes: no motility change during observation (pre-attached or mobile), transient attachment, permanent attachment and permanent detachment. For FimHF18 on 1M-BSA surfaces, 13.9% (251 out of 1,815 tracks) of all tracks showed a single transient attachment event (Supplementary Fig. 7d). In total, 67 out of the 251 bacteria that underwent a first transient adhesion attached and detached from the surface a second time. For these cells, the average time between detachment and re-attachment was only 13.5 s (Supplementary Fig. 7e), suggesting that re-binding may be favoured by proximity to the surface as compared with the initial attachment. The mean velocity on 1M-BSA, as compared with BSA-coated surfaces, was reduced for both FimHF18-piliated (4.2 and 7.4 μm s−1, respectively) and FimHF18–Ala188Asp-piliated bacteria (3.5 and 8.1 μm s−1, respectively; Supplementary Fig. 7a,b). This reduction of the mean velocity originates from two different phenomena: in FimHF18-piliated cells it is caused by a change from fast swimming to a slower mode of motion (Supplementary Fig. 7b; Supplementary Movie 1), which is consistent with bacterial surface rolling due to weak, short-lived mannose-based interactions29,53. In contrast, for FimHF18–Ala188Asp-piliated bacteria, the reduction of the mean velocity results from an increase in the fraction of adherent cells on 1M-BSA compared with BSA (see main text). In the presence of 200 μM HM, the mean velocity on 1M-BSA is increased for both FimHF18-piliated (6.5 μm s−1) and FimHF18–Ala188Asp-piliated bacteria (5.9 μm s−1; Supplementary Fig. 7a) and transient and permanent attachment is reduced by 75% and 85%, respectively (Fig. 6 and Supplementary Fig. 7c).
Accession codes: The atomic coordinates and structure factors have been deposited in the Protein Data Bank under the accession codes 4XO8, 4XO9, 4XOA, 4XOB, 4XOC, 4XOD, 4XOE.
How to cite this article: Sauer, M. M. et al. Catch-bond mechanism of the bacterial adhesin FimH. Nat. Commun. 7:10738 doi: 10.1038/ncomms10738 (2016).
Marshall, B. T. et al. Direct observation of catch bonds involving cell-adhesion molecules. Nature 423, 190–193 (2003).
Evans, E., Leung, A., Heinrich, V. & Zhu, C. Mechanical switching and coupling between two dissociation pathways in a P-selectin adhesion bond. Proc. Natl Acad. Sci. USA 101, 11281–11286 (2004).
Buckley, C. D. et al. Cell adhesion. The minimal cadherin-catenin complex binds to actin filaments under force. Science 346, 1254211 (2014).
Akiyoshi, B. et al. Tension directly stabilizes reconstituted kinetochore-microtubule attachments. Nature 468, 576–579 (2010).
Fiore, V. F., Ju, L., Chen, Y., Zhu, C. & Barker, T. H. Dynamic catch of a Thy-1-alpha5beta1+syndecan-4 trimolecular complex. Nat. Commun. 5, 4886 (2014).
Liu, B., Chen, W., Evavold, B. D. & Zhu, C. Accumulation of dynamic catch bonds between TCR and agonist peptide-MHC triggers T cell signaling. Cell 157, 357–368 (2014).
Das, D. K. et al. Force-dependent transition in the T-cell receptor beta-subunit allosterically regulates peptide discrimination and pMHC bond lifetime. Proc. Natl Acad. Sci. USA 112, 1517–1522 (2015).
Ronald, A. The etiology of urinary tract infection: traditional and emerging pathogens. Am. J. Med. 113, 14S–19S (2002).
Jones, C. H. et al. FimH adhesin of type 1 pili is assembled into a fibrillar tip structure in the Enterobacteriaceae. Proc. Natl Acad. Sci. USA 92, 2081–2085 (1995).
Hahn, E. et al. Exploring the 3D molecular architecture of Escherichia coli type 1 pili. J. Mol. Biol. 323, 845–857 (2002).
Waksman, G. & Hultgren, S. J. Structural biology of the chaperone-usher pathway of pilus biogenesis. Nat. Rev. Microbiol. 7, 765–774 (2009).
Thomas, W. E., Trintchina, E., Forero, M., Vogel, V. & Sokurenko, E. V. Bacterial adhesion to target cells enhanced by shear force. Cell 109, 913–923 (2002).
Zhou, G. et al. Uroplakin Ia is the urothelial receptor for uropathogenic Escherichia coli: evidence from in vitro FimH binding. J. Cell Sci. 114, 4095–4103 (2001).
Sharon, N. Carbohydrates as future anti-adhesion drugs for infectious diseases. Biochim. Biophys. Acta 1760, 527–537 (2006).
Ofek, I., Hasty, D. L. & Sharon, N. Anti-adhesion therapy of bacterial diseases: prospects and problems. FEMS Immunol. Med. Microbiol. 38, 181–191 (2003).
Guo, B. & Guilford, W. H. Mechanics of actomyosin bonds in different nucleotide states are tuned to muscle contraction. Proc. Natl Acad. Sci. USA 103, 9844–9849 (2006).
Le Trong, I. et al. Structural basis for mechanical force regulation of the adhesin FimH via finger trap-like beta sheet twisting. Cell 141, 645–655 (2010).
Han, Z. et al. Structure-based drug design and optimization of mannoside bacterial FimH antagonists. J. Med. Chem. 53, 4779–4792 (2010).
Wellens, A. et al. The tyrosine gate as a potential entropic lever in the receptor-binding site of the bacterial adhesin FimH. Biochemistry 51, 4790–4799 (2012).
Bouckaert, J. et al. Receptor binding studies disclose a novel class of high-affinity inhibitors of the Escherichia coli FimH adhesin. Mol. Microbiol. 55, 441–455 (2005).
Wellens, A. et al. Intervening with urinary tract infections using anti-adhesives based on the crystal structure of the FimH-oligomannose-3 complex. PLoS ONE 3, e2040 (2008).
Brument, S. et al. Thiazolylaminomannosides as potent antiadhesives of type 1 piliated Escherichia coli isolated from Crohn's disease patients. J. Med. Chem. (2013).
Vanwetswinkel, S. et al. Study of the structural and dynamic effects in the FimH adhesin upon alpha-d-heptyl mannose binding. J. Med. Chem. 57, 1416–1427 (2014).
Choudhury, D. et al. X-ray structure of the FimC-FimH chaperone-adhesin complex from uropathogenic Escherichia coli. Science 285, 1061–1066 (1999).
Schwartz, D. J. et al. Positively selected FimH residues enhance virulence during urinary tract infection by altering FimH conformation. Proc. Natl Acad. Sci. USA 110, 15530–15537 (2013).
Yakovenko, O. et al. FimH forms catch bonds that are enhanced by mechanical force due to allosteric regulation. J. Biol. Chem. 283, 11596–11605 (2008).
Vetsch, M., Sebbel, P. & Glockshuber, R. Chaperone-independent folding of type 1 pilus domains. J. Mol. Biol. 322, 827–840 (2002).
Gouin, S., Roos, G. & Bouckaert, J. Discovery and Application of FimH Antagonists 1–46Springer Berlin Heidelberg (2014).
Thomas, W. E., Nilsson, L. M., Forero, M., Sokurenko, E. V. & Vogel, V. Shear-dependent 'stick-and-roll' adhesion of type 1 fimbriated Escherichia coli. Mol. Microbiol. 53, 1545–1557 (2004).
Puorger, C. et al. Infinite kinetic stability against dissociation of supramolecular protein complexes through donor strand complementation. Structure 16, 631–642 (2008).
Thomas, W. et al. Catch-bond model derived from allostery explains force-activated bacterial adhesion. Biophys. J. 90, 753–764 (2006).
Lane, M. C. et al. Role of motility in the colonization of uropathogenic Escherichia coli in the urinary tract. Infect. Immun. 73, 7644–7656 (2005).
Wright, K. J., Seed, P. C. & Hultgren, S. J. Uropathogenic Escherichia coli flagella aid in efficient urinary tract colonization. Infect. Immun. 73, 7657–7668 (2005).
Nilsson, L. M., Thomas, W. E., Trintchina, E., Vogel, V. & Sokurenko, E. V. Catch bond-mediated adhesion without a shear threshold: trimannose versus monomannose interactions with the FimH adhesin of Escherichia coli. J. Biol. Chem. 281, 16656–16663 (2006).
Aprikian, P. et al. Interdomain interaction in the FimH adhesin of Escherichia coli regulates the affinity to mannose. J. Biol. Chem. 282, 23437–23446 (2007).
Petit, C. M., Zhang, J., Sapienza, P. J., Fuentes, E. J. & Lee, A. L. Hidden dynamic allostery in a PDZ domain. Proc. Natl Acad. Sci. USA 106, 18249–18254 (2009).
Tzeng, S. R. & Kalodimos, C. G. Protein dynamics and allostery: an NMR view. Curr. Opin. Struct. Biol. 21, 62–67 (2011).
Laskowski, R. A., Gerick, F. & Thornton, J. M. The structural basis of allosteric regulation in proteins. FEBS Lett. 583, 1692–1698 (2009).
Cooper, A. & Dryden, D. T. Allostery without conformational change. A plausible model. Eur. Biophys. J. 11, 103–109 (1984).
Tsai, C. J., del Sol, A. & Nussinov, R. Allostery: absence of a change in shape does not imply that allostery is not at play. J. Mol. Biol. 378, 1–11 (2008).
Pereverzev, Y. V., Prezhdo, O. V., Forero, M., Sokurenko, E. V. & Thomas, W. E. The two-pathway model for the catch-slip transition in biological adhesion. Biophys. J. 89, 1446–1454 (2005).
Pereverzev, Y. V. & Prezhdo, O. V. Force-induced deformations and stability of biological bonds. Phys. Rev. E Stat. Nonlin. Soft Matter Phys. 73, 050902 (2006).
Lou, J. Z. & Zhu, C. A structure-based sliding-rebinding mechanism for catch bonds. Biophys. J. 92, 1471–1485 (2007).
Manibog, K., Li, H., Rakshit, S. & Sivasankar, S. Resolving the molecular mechanism of cadherin catch bond formation. Nat. Commun. 5, 3941 (2014).
Choi, Y. I. et al. Dynamic control of beta1 integrin adhesion by the plexinD1-sema3E axis. Proc. Natl Acad. Sci. USA 111, 379–384 (2014).
Kong, F., Garcia, A. J., Mould, A. P., Humphries, M. J. & Zhu, C. Demonstration of catch bonds between an integrin and its ligand. J. Cell Biol. 185, 1275–1284 (2009).
Depoil, D. & Dustin, M. L. Force and affinity in ligand discrimination by the TCR. Trends Immunol. 35, 597–603 (2014).
Somers, W. S., Tang, J., Shaw, G. D. & Camphausen, R. T. Insights into the molecular basis of leukocyte tethering and rolling revealed by structures of P- and E-selectin bound to SLe(X) and PSGL-1. Cell 103, 467–479 (2000).
Preston, R. C. et al. E-selectin ligand complexes adopt an extended high-affinity conformation. J. Mol. Cell Biol. 8, 62–72 (2015).
Zakrisson, J., Wiklund, K., Axner, O. & Andersson, M. The shaft of the type 1 fimbriae regulates an external force to match the FimH catch bond. Biophys. J. 104, 2137–2148 (2013).
Whitfield, M. J., Luo, J. P. & Thomas, W. E. Yielding elastic tethers stabilize robust cell adhesion. PLoS Comput. Biol. 10, e1003971 (2014).
Xie, B. et al. Distinct glycan structures of uroplakins Ia and Ib: structural basis for the selective binding of FimH adhesin to uroplakin Ia. J. Biol. Chem. 281, 14644–14653 (2006).
Anderson, B. N. et al. Weak rolling adhesion enhances bacterial surface colonization. J. Bacteriol. 189, 1794–1802 (2007).
Pak, J., Pu, Y., Zhang, Z. T., Hasty, D. L. & Wu, X. R. Tamm-Horsfall protein binds to type 1 fimbriated Escherichia coli and prevents E. coli from binding to uroplakin Ia and Ib receptors. J. Biol. Chem. 276, 9924–9930 (2001).
Klein, T. et al. FimH antagonists for the oral treatment of urinary tract infections: from design and synthesis to in vitro and in vivo evaluation. J. Med. Chem. 53, 8627–8641 (2010).
Cusumano, C. K. et al. Treatment and prevention of urinary tract infection with orally active FimH inhibitors. Sci. Transl. Med. 3, 109ra115 (2011).
Sokurenko, E. V., Courtney, H. S., Ohman, D. E., Klemm, P. & Hasty, D. L. FimH family of type 1 fimbrial adhesins: functional heterogeneity due to minor sequence variations among fimH genes. J. Bacteriol. 176, 748–755 (1994).
Kleeb, S. et al. FimH antagonists: bioisosteres to improve the in vitro and in vivo PK/PD profile. J. Med. Chem. 58, 2221–2239 (2015).
Kuzmic, P. Program DYNAFIT for the analysis of enzyme kinetic data: application to HIV proteinase. Anal. Biochem. 237, 260–273 (1996).
Kabsch, W. Integration, scaling, space-group assignment and post-refinement. Acta Crystallogr. D Biol. Crystallogr. 66, 133–144 (2010).
McCoy, A. J. et al. Phaser crystallographic software. J. Appl. Crystallogr. 40, 658–674 (2007).
Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D Biol. Crystallogr. 60, 2126–2132 (2004).
Adams, P. D. et al. PHENIX: building new software for automated crystallographic structure determination. Acta Crystallogr. D Biol. Crystallogr. 58, 1948–1954 (2002).
Jo, S., Kim, T., Iyer, V. G. & Im, W. CHARMM-GUI: a web-based graphical user interface for CHARMM. J. Comput. Chem. 29, 1859–1865 (2008).
Phillips, J. C. et al. Scalable molecular dynamics with NAMD. J. Comput. Chem. 26, 1781–1802 (2005).
Essmann, U. et al. A smooth particle mesh Ewald method. J. Chem. Phys. 103, 8577–8593 (1995).
Sokurenko, E. V., Courtney, H. S., Maslow, J., Siitonen, A. & Hasty, D. L. Quantitative differences in adhesiveness of type 1 fimbriated Escherichia coli due to structural differences in fimH genes. J. Bacteriol. 177, 3680–3686 (1995).
Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).
We thank the staff at the Swiss Light Source (Villigen, Switzerland) for outstanding support for crystallographic data collection and the Imaging Facility as well as Research IT/A, and Mazur at Biozentrum, Basel, for supporting cell-tracking experiments. Discussions with Dr Jochen Reinstein (Max-Planck-Institute for Medical Research) are gratefully acknowledged. This work was supported by the Swiss National Science Foundation (Projects R’Equip 145023 (T.M) and Project 144183 (B.E., T.M. and R.G.), 310030B_138657 (R.G.) and 31003A_156304 (R.G.)). Calculations were performed at sciCORE (http://scicore.unibas.ch/) scientific computing core facility at University of Basel and the Swiss National Supercomputing Centre.
The authors declare no competing financial interests.
Supplementary Figures 1-7, Supplementary Note 1 and Supplementary References. (PDF 1837 kb)
Time-lapse analysis of FimHF18 (Abound) piliated E. coli cells on a mannosylated surface. Images were recorded over five minutes, segmented, background subtracted and converted into binary files by subsequent thresholding. The supplementary video is accelerated six-fold. In the first part (1‒16 s) raw images are shown, the second part (16‒26 s) is splitted into raw images (left) and binary images (right), the third part (26‒42 s) shows the processed images. (MOV 2594 kb)
Time-lapse analysis of FimHF18-Ala188Asp (Sbound state favored relative to FimHF18) piliated E. coli cells on a mannosylated surface. Images were recorded over five minutes, segmented, background subtracted and converted into binary files by subsequent thresholding. The supplementary video is accelerated six-fold. In the first part (1‒16 s) raw images are shown, the second part (16‒26 s) is splitted into raw images (left) and binary images (right), the third part (26‒42 s) shows the processed images. (MOV 1825 kb)
About this article
Cite this article
Sauer, M., Jakob, R., Eras, J. et al. Catch-bond mechanism of the bacterial adhesin FimH. Nat Commun 7, 10738 (2016). https://doi.org/10.1038/ncomms10738
This article is cited by
Nature Reviews Bioengineering (2023)
Biophysical Reviews (2023)
Nature Communications (2022)
Nature Communications (2022)
Optineurin links Hace1-dependent Rac ubiquitylation to integrin-mediated mechanotransduction to control bacterial invasion and cell division
Nature Communications (2022)