Abstract
The bacterial flagellum is a remarkable molecular motor, present at the surface of many bacteria, whose primary function is to allow motility through the rotation of a long filament protruding from the bacterial cell. A cap complex, consisting of an oligomeric assembly of the protein FliD, is localized at the tip of the flagellum, and is essential for filament assembly, as well as adherence to surfaces in some bacteria. However, the structure of the intact cap complex, and the molecular basis for its interaction with the filament, remains elusive. Here we report the cryo-EM structure of the Campylobacter jejuni cap complex. This structure reveals that FliD is pentameric, with the N-terminal region of the protomer forming an unexpected extensive set of contacts across several subunits, that contribute to FliD oligomerization. We also demonstrate that the native C. jejuni flagellum filament is 11-stranded and propose a molecular model for the filament-cap interaction.
Introduction
The bacterial flagellum is a macromolecular motor that rotates and acts as a propeller in many bacteria. It is associated with virulence in many human pathogens including Salmonella, enteropathogenic Escherichia coli, Campylobacter, and Helicobacter species 1,2. The flagellum is composed of > 25 different proteins, and consists of three main regions: the basal body acts as an anchor in the bacterial membrane, and includes the apparatuses for rotation and protein secretion; the hook forms a junction which protrudes from the outer membrane; and the filament, consisting of multiple repeats of a single protein (flagellin), forms the propeller 3. The filament, that can be > 20 μm in length, is topped by a cap complex, that consists of several copies of the protein FliD. This complex initially attaches to the hook-filament junction, and using a yet unknown mechanism, assists in building the filament 4.
Low-resolution cryo-EM studies of the cap complex in Salmonella enterica have suggested that it consists of five copies of FliD (also known as HAP2), forming a “stool”-shaped complex with a core “head” domain and five flexible “leg” domains, that interact with the growing end of the filament 5,6. Crystal structures of the FliD head domain have been reported for several species, and revealed a range of crystallographic symmetries, from tetramers in Serratia marscecens (FliDsm), pentamers in S. enterica (FliDse) and hexamers in E. coli (FliDec) and Pseudomonas. aeruginosa (FliDpa) 7–10. This observation led to the hypothesis that the cap complex can have a different, species-specific oligomeric states.
The flagellar filament has been studied extensively by cryo-EM, and its high-resolution structure has been reported in a range of bacteria, including Bacillus subtilis, P. aeruginosa and S. enterica. In all of these, the filament was shown to consist of 11 proto-filaments 11,12. However, a low-resolution cryo-EM study of the C. jejuni flagellar filament suggested the presence of 7 protofilaments 13. Taken together with the range of oligomeric states observed in the FliD crystal structures, these observations have led to a model where in different bacterial species, the cap complex has different oligomeric states (N), and in the corresponding filaments, the number of protofilaments is 2N + 1 7.
Campylobacter jejuni is a Gram-negative, spiral-shaped microaerophilic epsilon proteobacterium, colonizing the lower gastrointestinal (GI) tract of humans and poultry 14. It is often the most common cause of bacterial gastroenteritis and can lead to severe sequelae such as Guillain-Barré (GBS) and Miller-Fisher syndromes (MFS) 15. C. jejuni has two polar flagella located at each cell pole, which have an important function not only in motility, but are also responsible for adherence to surfaces, and for the secretion of virulence factor proteins 15,16. FliDcj is the major antigen in C.jejuni and thus a target for vaccine design 17–19.
In this study, we report the structure of the C. jejuni flagellar cap complex by cryo-EM. This structure demonstrates that FliDcj is pentameric, with an extensive set of contacts across several residues at the termini, that contribute to stabilizing the oligomeric state. We show that these interactions are essential for cell motility. We also observe that the full-length FliD protein for both S. marscecens (FliDsm) and P. aeruginosa (FliDpa) also form pentamers, with similar dimensions to that of FliDcj, indicating that the pentameric state of FliD within the cap complex is likely universal. Finally, we demonstrate that the native C. jejuni flagellum filament is 11-stranded, similar to other known flagellum filament structures. These observations allow us to propose a molecular model for the filament-cap interaction, and cap-mediated filament elongation.
Results
Cryo-EM structure of the flagellum cap complex
Existing high-resolution structures of FliD have so far been limited to the head domain. We therefore sought to characterize the intact FliD protein. To that end, we purified full-length FliD from several species: C. jejuni (FliDcj), P. aeruginosa (FliDpa) and S. marcescens (FliDsm) (Figure S1a). Size-exclusion chromatography demonstrated that all three proteins form oligomeric assemblies (not shown). However, preliminary negative-stain analysis showed that while the complexes formed by FliDpa and FliDsm are heterogeneous (Figure S1b), FliDcj forms homogeneous complexes, suitable for structural characterization.
Next we used cryo-EM to determine the structure of FliDcj. The protein forms discrete particles in vitreous ice, and 2D classification confirms that it adopts the dumbbell shaped structure previously reported for FliDst (Figure S2a). In addition, a significant subset of particles adopted top-view orientations, with clear 5-fold symmetry. This allowed us to obtain a structure of the full complex, to 4.71 Å resolution (Figures S2b, S2e).
The FliDcj complex possesses an overall architecture similar to FliDsm 5,6, consisting of ten subunits, with two pentamers interacting in a “tail-to-tail” orientation, through the leg domains (Figure 1a). A pentamer is about 170 Å in height (the decamer is ∼300 Å) and 130 Å in width with a 20 Å lumen (Figure 1c). We note however that the map shows a wide range of local resolution, with the leg domain well defined and with visible side-chains, while the head domain is much more poorly defined (Figure S2b).
This suggests that the complex is dynamic, with a hinge between the leg and head domains. To address this, we therefore performed a focused refinement on the head domain only, leading to a map at 5.02 Å resolution for this domain (Figure S2c, S2e). Using this map, we were able to generate an atomic model for this region of FliDcj, based on the crystal structure of FliDec (PDB ID: 5H5V) 8. We then used the map of the full complex to build the atomic model for the leg domain de novo (Figure S2d, S2f, Table 1).
The FliDcj structure shows that the FliD protomer folds in on itself in a ν-shape, which results in N and C termini next to each other in the leg domain. The overall architecture, as proposed previously, consists of a D0 domain formed by a long coiled coil, consisting of two helices located at the termini. A four-helix bundle forms the D1 domain. Connected to the D0-D1 leg domains are D2-D3 domains, rich in anti-parallel β-sheets, forming the head (Figure 1b) 7–9. This overall architecture is similar to that of the flagellin and hook, and in agreement with the previously reported structures of the FliD head domain 20. Intriguingly, while it was predicted that the D0 domain consists of a two-helix coiled-coil, as present in the flagellin and hook, our structure reveals that the N-terminal 17 residues are extended into a stretch that folds under and behind the monomer, interacting with the preceding subunit via a short β-strand. As a consequence, the C-terminal helix of the coiled-coil is not partnered with the N-terminus, but instead interacts with that of another molecule through hydrophobic interactions, forming the pentamer-to-pentamer interface. This intriguing architecture likely explains the strong tendency of FliD to form tail-to-tail complexes during isolation, as observed in FliDse 6 and FliDcj (this study).
We also note that FliDcj possesses a long insert within the D1 helix bundle, not present in other orthologues (Figure S3a). Secondary structure prediction indicates that this insert is likely globular (not shown), leading to the hypothesis that it forms an additional domain, termed D4. This type of domain insertion is not unusual, and has been observed in other FliD orthologues, as well as in flagellin and hook proteins 10,11,21,22. In our FliDcj map, we were able to observe density for this domain (Fig S3b), however it is at very low resolution, and did not allow us to build an atomic model. This suggests that the D4 domain is flexible. Indeed, further 3D classification revealed at least 4 distinct positions for this domain (Figure S3c). The role of this D4 domain is not known, but we postulate that it could be related to FliDcj’s capacity to bind to heparin, a feature involved in C. jejuni adherence but not observed in other FliD orthologues4.
Comparison with other FliD orthologues
Our cryo-EM structure of FliDcj is the first high-resolution structure of an intact FliD protein. Nonetheless the crystal structure of the head domain, corresponding to domains D2-D3, has been reported for a range of species, including S. enterica, P. aeruginosa, E. coli, S. marcescens, and H. pylori 6–10. In all orthologues, the structure is very similar, with RMSB values ranging from 1.5 Å to 2.5 Å to that of FliDcj (Figure 2a). In the E. coli orthologue, domain D1 was also present in the structure. It consists of a 4-helix bundle, and this structure is very similar to that of FliDcj, with a RMSD of 1.5 Å between the two structures. Nonetheless, we note that the position of D1 relative to that of D2-D3 is dramatically different in FliDec compared to FliDcj (Figure S4a). This suggests that the hinge between D1 and D2 is flexible, as supported by our focused refinement result.
In our cryo-EM map, FliD forms a pentameric architecture, consistent with the low-resolution cryo-EM structure of FliDse, with a similar overall architecture consisting of two pentamers in a head-to-tail arrangement. In contrast, crystal structures of the head domains from FliD in several species reported a range of oligomeric states, including tetramer (FliDsm), pentamer (FliDse) and hexamers (FliDpa and FliDec)7–9. When comparing the dimensions of these structures, the diameters of all complexes are similar, around ∼140 Å. However, the dimension of the lumen differs significantly between structures, with FliDcj and FliDse having a central lumen of ∼20 Å, while FliDpa and FliDec have a lumen of ∼50 Å and ∼40 Å respectively, and FliDsm a ∼15 Å lumen (Figure 2b). Even in the case of FliDse, which crystallized as a pentamer, while the overall dimensions are similar to that of the head domains of the FliDcj pentamer, in the E. coli orthologue the pentamer is flattened compared to that of FliDcj (Figure S4b). Based on our structure, we hypothesize that there is a large degree of plasticity in the interface between the D2-D3 domains of adjacent molecules, and therefore in the absence of D0, a range of interfaces can be trapped in the crystal contacts. We propose that the additional contacts formed by the N-terminal stretch are essential for FliD to adopt its true oligomeric state.
To verify this, we investigated the oligomeric state of full-length FliDsm and FliDpa, the head domains of which crystallized as tetramers and hexamers, respectively, by negative stain TEM. As mentioned above, these proteins do not form uniform complexes (Figure S1b). Nonetheless, we noted that the majority of the particles appeared as top views, which allowed us to perform preliminary 2D classification to determine their lateral symmetry. This revealed that both orthologues form pentamers with similar dimensions to that of FliDcj (Figure 2c). However, in the FliDpa sample we observed additional particles with 6-fold and 4-fold symmetry, while in the FliDsm sample there was a large percentage of particles with 4-fold symmetry. The dimensions of the particles in those 2D classes are significantly larger than the FliD pentamer, and therefore we could not conclude if these correspond to alternative oligomeric species, or to other negative stain artifacts and/or non-specific aggregates. However, the presence of pentamers with similar dimensions to that of FliD supports the hypothesis that the native architecture of the cap complex is a FliD pentamer, with contacts at the N-terminus required for FliD to adopt its true oligomeric state.
Hydrophobic interactions in the D0 domain are required for forming functional filaments
As mentioned above, our structural characterization of the cap complex indicates an unusual architecture of the N-terminus, which forms a stretch that wraps around and forms contacts with two adjacent subunits, through hydrophobic contacts (Figure 3a). In particular, Residues Leu 9 and Phe 11 are buried within a pocket formed by Trp 614 and Tyr 617, located in the C-terminus of the adjacent molecule. This is of particular interest since it was shown that the C-terminus contributes to the oligomerization of FliD and interaction with its chaperone 23. We also note that both the N- and C-termini of FliD are highly conserved across species, with mainly aromatic side-chains present in all orthologues in the aforementioned positions.
To confirm the role of these residues in FliD function, we engineered a C. jejuni fliD knockout strain (ΔfliD), leading to a loss of motility in a soft agar swarm assay. Accordingly, no filament was observed in this strain (Figure S5a). Genetic complementation by expressing the fliD gene at a distal site on the chromosome fully rescued motility (Figure 3b, S5a), and we exploited this to engineer point mutations in the aforementioned residues to assess their impact on motility.
Mutation of Leu 9, Phe 11, Trp 614 or Trp 617 significantly reduced motility, up to 40% for mutations to polar residues (F11S, W614S, L9S or Y617S) (Figure 3b). This confirms that the hydrophobic properties of these residues are critical for motility, suggesting that the interaction formed by the N-terminal stretch contributes to FliD function. To verify if motility was affected because the aforementioned mutations prevented filament assembly, we visualized the corresponding bacteria by TEM. All of the mutations still led to bacteria with assembled filaments, of length similar to that of WT bacteria (Figure S5b), demonstrating that the corresponding FliD proteins are still able to promote filament elongation. However, we noted that the filaments are much more brittle in the mutants, with between 60 and 80% of filaments found unattached to the bacterial cell, versus ∼ 20% in the WT bacteria (Figure S5c). We also note that the N-terminal ∼ 20 residue stretch corresponds to the secretion signal in flagellar filaments of S. enterica, so potentially a similar signal exists for FliD to be secreted through the flagellum T3SS 24. The observation that in the mutants described above, the filament is still formed, is a strong confirmation that these mutations did not interfere with FliD secretion, but rather with its function to promote filament elongation. The second set of interactions observed in the D0 domain, is formed between the C-terminus of FliD in the pentamer-to-pentamer interface (Figure 3a). Evidence from tomography, as well as other biochemical data, indicate that this interaction is not physiological 5,25–27. However, since it is observed in both FliDcj and FliDse, we postulated that it mimics the interaction between FliD and the filament. To verify this, we engineered a series of mutations in the residues forming this interface (Leu 628, Ile 635, Leu 624 and Ile 620) and characterized their impact on motility as described above (Figure 3b). Mutating these residues impacted motility, however the effect is less pronounced than the mutants involved in the N-terminal stretch interaction, with the exception of I620S and L624S mutations. We propose that this is because the overall hydrophobic propensity, rather than specific contribution of each amino acid, is the critical element of this region of the protein.
A structural model of the C. jejuni filament
Current Cryo-EM structures of various flagellar filaments have demonstrated that they consist of 11 protofilaments, formed by a single protein, the flagellin 11. The flagellin consists of four domains D0-D3, and can adopt two conformations, termed L and R, leading to two alternative filament structures, left-handed and right-handed, respectively. C. jejuni possesses two flagellin homologues, FlaA and FlaB, that are ∼ 95% identical to each other, with both required for the formation of fully functional filaments. FlaA and FlaB are highly similar to other flagellins (Figure S6a), except for an ∼ 70 amino acid insert in D2 that likely consists of a globular insert, as observed in several flagellin orthologues 11. Surprisingly, a previously published EM structure of the C.jejuni filament had reported a 7 protofilament arrangement 13. However, this structure was obtained from a FlaA G508A mutant, in the absence of FlaB, and is at low resolution. It is therefore not clear if this was an artifact and/or wrong interpretation of the data, or if the C. jejuni filament indeed possesses a different architecture to other species.
To reconcile this, we sought to determine the structure of the native C. jejuni filament, directly from wild type cells (Figure 4a). To avoid biases due to symmetry, we initially performed a reconstruction without any helical symmetry applied. This map clearly possessed 11-fold symmetry (Figure S6b), despite the low resolution (∼ 27 Å, figure S6c). This demonstrates that the C. jejuni flagellar filament consists of 11 protofilaments with a lumen of ∼25-30 Å and outer diameter of ∼200 Å, similar to that of other bacterial species. We therefore refined the map further by applying helical symmetry, with a 65.4° twist and 7.25 Å rise, which allowed us to reach ∼ 8.6 Å resolution (Figure S6c). In this map the central D0-D1 domains are well resolved, with the density for helices clearly visible (Figure 4b). The density for domains D2 and D3 is visible, but less well resolved. The fact that we can only reach limited resolution is perhaps not surprising, since we likely have a combination of L and R conformations for the flagellin.
Based on this, we used the previously published structure of the P. aeruginosa filament (PDB ID: 5WK6), to position the cap complex within the filament structure. This allowed us to propose a model for FliD-flagellin interaction (Figure 4c). In this model, the C-terminus of FliD forms broadly non-specific, hydrophobic contacts with exposed regions of the filament, similar to flagellin-flagellin interactions (Figures 4c and 4d). A gap between adjacent FliD molecules, on the side of the leg domain, is positioned in a suitable location for the insertion of a flagellin molecule and is the likely site of exit for nascent molecules (Figure 4d). This however remains to be verified experimentally.
Discussion
In previous studies, evidence suggesting different stoichiometries for the flagellum filament and/or cap complex in different species was based on low-resolution cryo-EM structures, and crystallographic symmetries of truncated proteins. Here we largely resolve this conflicting evidence, by demonstrating that FliD adopts a pentameric stoichiometry in a range of species, and that the filament of C. jejuni is 11-stranded, and not 7-stranded as reported previously. We can therefore conclude that the stoichiometry of these proteins is conserved across species, with a 11-to-5 asymmetry between these two different regions of the bacterial flagellum. Our structure of the intact cap complex, supported by mutagenesis studies, suggests that the FliD C-terminal domain interaction with the opposite pentamer in the decametric complex mimics that of the FliD interaction with the filament. We hypothesize that exposed hydrophobic residues, both on the D0 domain of flagellin molecules and in the C-terminus of FliD, act as a chaperonin-like environment to promote the folding and insertion of new flagellins 28,29.
Based on the results reported in this study, we propose a universal mechanism for cap-mediated filament elongation, as illustrated in figure 5. The FliD cap pentamer fits into the flagellum filament, through interactions between D0 of flagellins and the C-terminus of FliD. Because of the symmetry mismatch, this interaction is not present on one side of the cap complex. New flagellin molecules are secreted through the filament, and ultimately enter a chamber inside the cap complex (1). The flagellin then exits this cavity through a lateral opening, where the location of the next flagellin insertion site is positioned (2). The four other cavities are sterically blocked by the flagellum filament. There, exposed hydrophobic residues act as a chaperone, and promote flagellin folding in its insertion site (3). The folding of the new flagellin protomer leads to dislodging of the cap complex, that rotates by ∼ 35 ° (4), thus positioning an adjacent cavity of the cap complex close to the next flagellin insertion site (Figure 5).
We note that previous studies, based on low-resolution tomography data, have suggested that the D0 domain of FliD might be dynamic, with the leg domains opening and closing to promote filament elongation 5,9,26. Our structure of the cap complex does not support this model, as we show that the N-terminal stretch of FliD is essential for filament elongation and maintains the leg domains in a rigid position. Our data supports an alternative mechanism, which had been proposed previously, whereby the cap complex acts as a rigid cog that rotates during flagellum elongation 8. Further experiments to characterize the flagellum-cap complex at high resolution will be required to confirm this model.
In conclusion, we report the cryo-EM structure of the flagellum cap complex, and demonstrate that FliD across multiple species (FliDsm, FliDpa and FliDse) forms pentameric complexes. We show that the interface between opposite D0 leg domains in the FliD decamer complex is essential for cell motility and formation of a functional filament, and therefore likely plays a role in FliD-filament interactions. We also demonstrate that the C. jejuni flagellar filament possesses the same architecture as that of other species. Taken together, these results allow us to propose a universal model for cap-filament interaction as well as propose a mechanism for cap-mediated filament elongation.
Materials and Methods
Protein Expression and purification
The genes coding for FliDcj, FliDsm and FliDpa, codon-optimized for expression in E. coli, were synthesized (BioBasic) and sub-cloned into pET28a (Novagen). Recombinant proteins were expressed in E. coli BL21-CodonPlus(DE3)-RIL cells containing the corresponding plasmids. For FliDcj, transformants were grown in LB medium at 37 °C until they reached log phase, and expression was induced by the addition of 1mM IPTG overnight at 20 °C. For both FliDpa and FliDsm, expression was auto-induced in ZYM-505230 media at 20 °C overnight. For all three proteins, cells were collected by centrifugation, resuspended in 50 mM HEPES 150 mM NaCl pH 7 and sonicated. The lysate was centrifuged at 14 000g at 4 °C for 45 minutes. The supernatants were applied onto a 5 ml HisPure™ Ni-NTA resin (ThermoScientific) gravity-based column equilibrated with 50 mM HEPES 150 mM NaCl pH 7 and eluted using a linear 20-500 mM Imidazole gradient. Fractions containing FliD were pooled and applied to a HiLoad Superdex 200 16/600 column (GE Healthcare) equilibrated with 50 mM HEPES 150 mM NaCl pH 7 for FliDcj and 50 mM Tris 150 mM NaCl pH 8 for FliDpa and FliDsm.
Negative-stain grid preparation and data collection
For negative-stain TEM experiments, ∼ 5 μl of purified protein, or of cell culture in log phase, was applied onto glow-discharged, carbon-coated copper grids (Agar Scientific). After incubating the sample for ∼2 minutes at room temperature, the grids were rapidly washed in three successive drops of deionized water and then exposed to three successive drops of 0.75% uranyl formate solution. Images were recorded on a CM100 TEM (Phillips) equipped with a MSC 794 camera (Gatan) (FliDcj and C. jejuni cell cultures) or a Technai T12 Spirit TEM (Thermo Fisher) equipped with an Orius SC-1000 camera (Gatan). Datasets were manually acquired with a pixel size of 2.46 Å/pix, and a defocus range from −0.8 μm to −2.0 μm. The micrographs were processed using cisTEM 31 package, with CTF parameters determined by CTFFIND4 32. Approximately 3500 particles were picked for FliDsm and 2700 for FliDpa to generate representative two-dimensional (2D) class averages with 330 Å mask diameter. The point mutant flagella attachment was determined through imaging grids at 700x magnification at about 20 micrographs per mutant containing cell count from 30 to 100 cells. The percentage of attachment was calculated as a proportion of the total flagella observed per mutant.
Cryo-EM grid preparation and data collection
For the structural characterization of FliDcj, aliquots of (5 μl) of purified protein at a concentration of 1 mg ml-1 was deposited onto glow-discharged C-flat holey carbon films 1.2/1.3 200 mesh (EMS). A Vitrobot Mark III (FEI) plunge-freezing device was used for freeze-plunging, using double-blotting 33 with a final blotting time of 6.5 seconds. Cryo-EM data were collected with a Titan Krios TEM operated at 300 kV and equipped with an energy filter (Gatan GIF Quantum) and recorded on a K2 Summit direct electron detector (Gatan) operated in counting mode. 1223 micrographs were automatically acquired with the EPU software (Thermo Fisher), at a pixel size of 1.38 /pix, using a total dose of 41 e- Å-2 and with 40 frames per micrograph. The defocus range used for data collection was −1.0 μm to −2.6 μm.
For the structural characterization of the native C. jejuni filament, wild-type 81116 strain cell culture grown to OD600 = 5 was applied onto glow-discharged C-flat holey carbon films 2/2 200 mesh (EMS). A Leica EM GP (Leica) plunge-freezing device was used for freezing, with a 6 s blotting time. Cryo-EM data were collected on a Technai Arctica TEM (Thermo Fisher) operated at 200 kV and equipped with a Falcon III camera. 100 micrographs were collected using the EPU software (Thermo Fisher) in linear mode, with a pixel size of 2.03 Å/pix, with a total dose of 45 e- Å-2 and 1 frame per micrograph. The defocus range used for data collection was approximately −0.8μm to −2.0 μm.
Cryo-EM image processing and reconstruction
For FliDcj, processing was done in RELION 2.0 34. Motion correction was performed with MotionCor2 35, with dose-weighting. CTF parameters were determined by CTFFIND4 32 software. Approximately 2000 particles were manually picked from selected micrographs to generate representative 2D class averages. These classes were used as templates for automated particle picking for the entire dataset. A total of 130000 particles were picked and extracted using a 280 × 280 pixels box. After multiple rounds of 2D classification, 55967 particles from the best 2D classes were obtained and used to generate an initial model. Following further 3D classification and refinement with D5 symmetry, a final map to 4.71 Å resolution was generated, which was sharpened using PHENIX 1.13 36. The leg domains were visibly at a higher resolution than the head domains, therefore a mask centering on the head domain was used for further refinement with C5 symmetry, leading to a map of the head domain to 5.02 Å resolution. Further 3D classification of the masked head domain was used to identify 4 different conformations of the D4 domain not resolved in the full map. For the native C. jejuni filament, processing was done in RELION 3.0 34. Motion correction was performed with MotionCor2 35, with dose-weighting. CTF parameters were determined by CTFFIND4 32. Filaments were manually picked, and particles were extracted using a 7.6 Å rise and 300 pixel box leading to a set of 254041 segments. Multiple rounds of 2D classification gave a final dataset of 71828 good particles which were used for 3D refinement, both with and without imposed helical symmetry. Without symmetry, the structure refined to 27.2 Å resolution, but when helical symmetry was applied, the final resolution after further classification and refinement was 8.6 Å, with a 65.4° twist and a 7.25 Å rise.
Model building and refinement
For the D2-D3 domains, a homology model was generated with PHYRE2 37, using the FliDec crystal structure 8 (PDB:5H5V) as a template. These domains were fitted into the sharpened map in Chimera 38. This model was subjected to iterative rounds of real-space refinement and building using PHENIX 1.16 36 and Coot39 respectively. The N-terminal stretch was modeled with RosettaES 40, and then the remaining missing loops were modeled using RosettaCM41 guided by the electron density. The output model was refined once more in Coot to improve the geometry and delete any modelled residues in areas without electron density.
Cultivation of C. jejuni
C. jejuni strain 81116 was grown on blood agar plates (Colombia base agar with 5% v/v defibrinated horse blood) in a microaerobic cabinet (Don Whitley, UK) at 42°C with a controlled atmosphere of 10% v/v O2, 5% v/v CO2 and 85% v/v N2. Where appropriate, the selective antibiotics kanamycin and chloramphenicol were added at 50 µg/ml and 20 µg/ml, respectively.
Construction of fliD deletion mutant and complemented strains
A fliD mutation vector was constructed using NEB HiFi DNA assembly method (E2621, New England Biolabs). Briefly, flanking regions of fliD were amplified from C. jejuni 81116 genomic DNA using primers fliDmutantF1-R2 (Table S1). These flanks were assembled into pGEM3ZF either side of a non-polar kanamycin resistance cassette, amplified from pJMK30 using primers KanF/R (Table S1). The final mutation vector was designed such that spontaneous double crossover with the C. jejuni 81116 genome would result in the replacement of the majority of the open reading frame of fliD with the kanamycin resistance cassette, allowing a means of selection. For complementation of the mutant, fliD was amplified from C. jejuni 81116 genomic DNA using primers fliDcompF/R (Table S1). The amplified fragment was digested with BsmBI at sites incorporated into the primers and ligated into similarly digested pCmetK plasmid, a complementation vector for C. jejuni incorporating flanking regions of the pseudo-gene region corresponding to cj0046 in C. jejuni 11168 to allow insertion into the genome, a constitutive promoter from the C. jejuni metK gene to drive expression of fliD, and a chloramphenicol resistance cassette. To generate the strains, wildtype C. jejuni 81116 was first transformed with the fliD mutation vector by electroporation and colonies selected for kanamycin resistance on blood agar plates. The isolated mutant strain was then further transformed with the fliD complementation vector and selected for double kanamycin / chloramphenicol resistance.
Construction of fliD point mutants in C.jejuni
Point mutations in fliD were constructed by site directed mutagenesis of the complementation vector using the KLD method (M0554, New England Biolabs). Briefly, the fliD complementation plasmid was amplified by PCR with divergent primers containing targeted nucleotide substitutions in the forward primer (listed in Table S2). An aliquot of the linear PCR product was treated with the KLD enzyme mix to circularise the mutated plasmid while degrading any residual template. The treated plasmids were transformed into E. coli DH5α and transformants selected by chloramphenicol resistance. Plasmid was purified from multiple transformants and the fliD open reading frame was sequenced to ensure the correct substitution had been introduced without secondary mutations (LightRun sequencing, Eurofins EU). Point mutated complementation vectors were then transformed into the C. jejuni fliD mutant strain as above to generate the collection of point mutant strains.
Motility assays
Overnight growth of C. jejuni on blood agar plates was harvested and resuspended in phosphate buffered saline to an optical density at 600 nm of 1.0. 0.5 µl aliquots were then injected into semi-solid agar plates (0.4 % w/v agar, 3.7 % w/v brain heart infusion) containing 5×10−3 % triphenyl tetrazolium chloride, a redox dye which allows clear visual assessment of growth. The diameter of growth was measured after 16 hours of incubation.
Data availability
The map for FliDcj is available at EMDB with accession code EMD-10210, and the atomic model is available in Protein Data Bank with accession code 6SIH. The map for the native filament is available at EMDB with accession code EMD-10244. All other data supporting the findings of this study are available from the corresponding authors upon request.
Author contributions
N.S.A. and J.R.C.B. conceived the project and designed the structural experiments. A.J.T. and D.J.K. designed the C. jejuni cloning, mutagenesis and motility assays. N.S.A. performed the protein purification, Cryo-EM data collection and processing, as well as the negative stain experiments. A.J.T. performed the C. jejuni mutagenesis and motility assays, together with N.S.A. S.T. provided assistance with data collection and setup of electron microscopy facility. D.F. and F.D. refined the FliDcj atomic model with Rosetta. All authors contributed to the writing and editing of the manuscript.
Competing interests
The authors declare no competing interests
Acknowledgements
This work was funded by a UK Biotechnology and Biological Sciences Research Council (BBSRC) grant (BB/R009759/1) to J.R.C.B. N.S.A. was recipient of PhD scholarship from The Global Strategic Alliance at the University of Sheffield. A.J.T. was funded by a BBSRC grant (BB/R003491/1) to D.J.K. We thank the members of EM facility for their essential assistance and microscope access, and we acknowledge the members of Prof. Per Bullough’s laboratory for fruitful discussions. Cryo-EM data for FliDcj was collected at the UK national Electron Bio-Imaging centre (eBIC), proposal EM19709-1. The C.jejuni filament data was collected at the University of Sheffield Electron Microscopy Facility.
Footnotes
↵* d.kelly{at}sheffield.ac.uk