Significance Many bacteria swim using helical propellers, flagella. Intriguingly, different bacteria show different swimming abilities, strikingly illustrated by the abilities of some to bore through viscous fluids (e.g., gastrointestinal mucus) in which others are completely immobilized. We used 3D electron microscopy to show that differences can be explained by the structures of the torque-generating motors: two diverse high-torque motors position additional torque-generating complexes at wider radii from the axial driveshaft than in the model enteric bacteria; this positioning is consistent with the exertion of greater leverage to rotate the flagellum and thus greater torque generation. Intriguingly, these torque-generating complexes are scaffolded at wider radii by a conserved but divergent family of structures, suggesting an ancient origin of reconfiguring torque output.

Abstract Although it is known that diverse bacterial flagellar motors produce different torques, the mechanism underlying torque variation is unknown. To understand this difference better, we combined genetic analyses with electron cryo-tomography subtomogram averaging to determine in situ structures of flagellar motors that produce different torques, from Campylobacter and Vibrio species. For the first time, to our knowledge, our results unambiguously locate the torque-generating stator complexes and show that diverse high-torque motors use variants of an ancestrally related family of structures to scaffold incorporation of additional stator complexes at wider radii from the axial driveshaft than in the model enteric motor. We identify the protein components of these additional scaffold structures and elucidate their sequential assembly, demonstrating that they are required for stator-complex incorporation. These proteins are widespread, suggesting that different bacteria have tailored torques to specific environments by scaffolding alternative stator placement and number. Our results quantitatively account for different motor torques, complete the assignment of the locations of the major flagellar components, and provide crucial constraints for understanding mechanisms of torque generation and the evolution of multiprotein complexes.

Flagellated bacteria have tailored their motility to diverse habitats. For example, the enteric model organisms Salmonella enterica serovar Typhimurium and Escherichia coli colonize animal digestive tracts and can reside outside a host, assembling flagella over their cell body to swim. However, a diverse spectrum of flagellar swimming ability is seen across the bacterial kingdom. Caulobacter crescentus inhabits low-nutrient freshwater environments where it swims using a high-efficiency flagellar motor (1, 2), whereas Vibrio species produce high-speed, sodium-driven polar flagella to capitalize on the high sodium gradient of their marine habitat (3). On the other hand, the ε-proteobacteria and spirochetes, many of which thrive exclusively in association with a host, have evolved characteristically rapid and powerful swimming capabilities that enable them to bore through mucous layers coating epithelial cells or between tissues. Indeed, the ε-proteobacteria Campylobacter jejuni and Helicobacter pylori are capable of continued swimming in high-viscosity media that immobilize E. coli or Vibrio cells (4⇓–6), and similar behavior is observed for spirochetes (7, 8).

Despite differences in the organisms’ swimming ability, the flagellar motor is composed of a conserved core of ∼20 structural proteins (9). The mechanism of flagellar motility is conserved (10), with torque generated by rotor and stator components (9). Stator complexes, heterooligomers of four motility A (MotA) and two motility B (MotB) proteins, are thought to form a ring that surrounds the axial driveshaft. Transmembrane helices of MotA and MotB form an ion channel, and MotB features a large periplasmic domain that binds peptidoglycan (11, 12) and the flagellar structural component, the P-ring (13). The stator complex couples ion flux to exertion of force on the cytoplasmic rotor ring (the C-ring), which transmits torque to the axial driveshaft (the rod), universal joint (the hook), and helical propeller (the filament), culminating in propulsion of the bacterium. Biophysical (14) and freeze-fracture (15) studies together with modeling (16) have proposed that a tight ring of ∼11 stator complexes dynamically assembles around the rod above the outer lobe of the C-ring in closely related Salmonella and E. coli motors (which we collectively refer to as the “enteric motor”). However, despite these conclusions, and although the structures observed in subtomogram averages have been proposed to be the stator complexes (17⇓–19), the locations and stoichiometries of the stator complexes remain to be confirmed.

How can we explain the wide diversity in flagellar swimming abilities in the context of a conserved core flagellar motor? Biophysical studies suggest that the source of the difference lies, at least in part, in variations in the mechanical output of the motors themselves. Torques of motors from different bacteria have been shown to range over an order of magnitude, and torque correlates with swimming speed and the ability of bacteria to propel themselves through different viscosities, indicating that adaptations are likely to be at the level of the motor itself. [Torque also varies within a single species, up to a maximum value, as a function of the number of stator complexes incorporated into the motor (14)]. For example, C. crescentus motors have been measured to produce torques of 350 pN⋅nm (2). Estimates for the torque of the enteric motor ranges from ∼1,300 to ∼2,000 pN⋅nm (20, 21). The ε-proteobacterium H. pylori has been estimated to swim with torque of 3,600 pN⋅nm (22), and spirochetes are capable of swimming with 4,000 pN⋅nm of torque (21, 23). Sodium-driven motor torques in Vibrio spp. have been measured between ∼2,000 and 4,000 pN⋅nm (24), depending on the magnitude of the sodium gradient. It is noteworthy, however, that an estimated sodium motive force in Vibrio spp. that is lower than the standard E. coli proton motive force nevertheless drives the Vibrio motor with higher torque than the E. coli motor (24, 25), further suggesting that torque differences likely exist at the level of the motor. However, the molecular mechanism by which different motors might produce different torques has not been investigated.

The simplest scenario for tuning motor torque would be evolved adaptation of motor architecture. In support of this scenario, we recently showed that many motors have evolved additional structures not found in the well-studied enteric motors (18), and we observed that the C-ring radius varies among species (17, 18). One of the most widespread novel structures is a periplasmic basal disk directly beneath the outer membrane, often co-occurring with varied uncharacterized additional structures, which we collectively term “disk complexes.” Consistently, disk complexes have been seen only in motors that produce torque higher than that in E. coli or Salmonella. For example, the sodium-driven ∼2,000+ pN⋅nm torque motors of Vibrio species assemble a disk complex featuring a basal disk beneath the outer membrane (18) in addition to smaller H- and T-rings composed of FlgOT (flagella O, T) and MotXY (motility X, Y), respectively (26, 27). It has been shown that the T-ring interacts with stator complexes in Vibrio spp. (28), although the exact location and number of stator complexes in Vibrio spp. remains unclear. ε-Proteobacteria such as Helicobacter species, C. jejuni, and Wolinella succinogenes also assemble disk complexes composed of large basal disks beneath the outer membrane together with additional smaller disks (18, 29). Although these and other cases of additional disks have been reported (18, 30), their relation to flagellar function remains enigmatic, and it is unclear if these widespread disk complexes are homologous or analogous.

In this study, we hypothesized that bacteria have tuned their swimming abilities by evolving structural adaptations to their flagellar motors that would result in altered torque generation. Using electron cryo-tomography and subtomogram averaging, we found that Vibrio polar γ-proteobacterial and Campylobacter ε-proteobacterial flagellar motors incorporate 13 and 17 stator complexes, respectively, compared with the ∼11 in enteric bacteria. In both cases, these stator complexes are scaffolded into wider stator rings relative to the enteric motor by components of their respective disk complexes. The wider C. jejuni stator ring is further reflected in a considerably wider rotor C-ring. Further analysis of the components of the Vibrio and C. jejuni disk complexes reveals that they share a core protein, FlgP, but each has acquired diverse additional components to form divergent disk-complex architectures. We conclude by showing that our structural data of wider stator rings featuring additional stator complexes can quantitatively account for the differences in torque between different flagellar motors.

Discussion In this work we sought to determine whether the generation of high torque by the flagellar motors of specific bacterial species could be functionally attributed to the unique disk complexes associated with their respective motors. For this approach, we analyzed the flagellar motors of two bacterial species, V. fischeri and C. jejuni, with a combination of genetic and biochemical analysis of bacterial mutants and electron cryo-tomographic imaging and compared these motors with the enteric motor assembled in Salmonella. Our studies indicate that the disk complexes of these two organisms, although displaying structural diversity and composed of both homologous and unrelated proteins, share a common role in forming a platform for the integration of different numbers of stator complexes into the flagellar motor. Furthermore, these disk complexes not only integrate more stator complexes into a motor but also place them at wider radii from the center of the motor’s axis of rotation than seen in the enteric motor. As discussed in detail below, we propose that this creation of a wider stator ring with an increased number of stator complexes inevitably results in increased torque in these motors, directly affecting the swimming ability of the respective bacteria by conferring higher swimming velocities and the ability to migrate through environments with increased viscosity. A Proposed Biogenesis Order for Assembly of the C. jejuni Disk Complex. Although parts of the purified Vibrio motor have been studied previously, no components of the C. jejuni disk complex had been identified. Identifying the components of the C. jejuni motor and examining the structures of mutants of these components enable us to propose an assembly pathway for the biogenesis of a functional C. jejuni flagellar motor (Fig. 4B). This biogenesis pathway is likely divided into two stages. The first stage is the formation of a nonfunctional C. jejuni core flagellar motor that resembles the (functional) Salmonella motor, as evidenced by the nonfunctional ΔflgP or ΔflgQ C. jejuni motors that fail to incorporate any disk-complex components (Fig. 4A). The second stage involves the sequential assembly of each disk in the disk complex, ultimately enabling correct incorporation of the stator complexes to produce a functional motor. Consistent with our findings that FlgP requires core flagellar components for outer membrane association, we suspect that FlgP polymerizes into the basal disk (with assistance from FlgQ) only after the core flagellar motor forms to provide an attachment site for the basal disk (possibly at the P- and/or L-ring). Besides FlgQ, no other disk-complex proteins are required for FlgP to localize to the outer membrane or form the basal disk (Figs. 4 and 5 A and B). Formation of the basal disk provides a platform and site of interaction with PflA to form the inner node of the medial disk (Fig. 4). As the medial disk subsequently forms, PflA serves as a platform for PflB together with the stator complexes to assemble into the proximal disk (Fig. 4). The disk complex therefore forms a separate and previously unappreciated additional biogenesis step in ε-proteobacteria that occurs after the conserved assembly of the flagellar type III secretion system, rotor, switch, rod, hook, and filament. FlgP-Based Disk Complexes Are Widespread. Because C. jejuni and V. fischeri FlgP proteins share low sequence identity, FlgP may represent an ancient recruitment to the ancestral motor, followed by later independent evolutionary recruitment of the remaining Vibrio or ε-proteobacterial disk proteins to create motors with disk complexes that incorporate additional stator units into the motor. The speculation that FlgP recruitment was the first step in the evolution of disk complexes is supported further by the morphological divergence and mutually exclusive sets of additional accessory proteins between the two structural variants. Widespread presence of FlgP and other C. jejuni disk-complex proteins (i.e., FlgQ, PflA, and PflB) across the ε-proteobacteria, together with previous observations of flagellar motors incorporating disk complexes (18, 29), confirm that other ε-proteobacteria such as Helicobacter and Wolinella genera also assemble C. jejuni-type homologous disk complexes. FlgP also is widespread among many γ-proteobacterial genera including Vibrio, Shewanella, and Rhodobacter, but in these genera components of the Vibrio disk complex MotX, MotY, FlgO, and FlgT are encoded, as is consistent with assembly of Vibrio-type disk-complex homologs (45, 46). Intriguingly, however, the widespread FlgP basal disks do not interact directly with the stator complexes in either the Vibrio or ε-proteobacterial disk-complex variants. In V. fischeri, FlgP connects the T-ring to the outer membrane, and components of the H-ring interact with the stator complexes. In the absence of FlgP and the basal disk, the T- and H-rings are produced, but we suspect that their architecture may be altered so that these rings cannot interact with a stator complex. In C. jejuni, the FlgP basal disk is required to assemble the remainder of the disk complex, with the proximal disk responsible for incorporation of stator complexes. Nevertheless, the common role of FlgP in ultimately creating scaffolds for stator-complex incorporation in both C. jejuni and V. fischeri suggests that their common ancestor had the same function of providing scaffolding for the stator complex, a function that was retained during divergence of the two forms of disk complexes by the incorporation of the additional proteins. Alternatively the two disk-complex lineages may have independently evolved roles to scaffold additional stator complexes. We now are pursuing a follow-up study to understand the phylogeny and probe the origins of these disk complexes. Interactions of Stator Complexes with Flagellar Motors Vary in Motile Bacterial Species. The interactions of stator complexes with the disk complexes of the Vibrio and Campylobacter motors are different from those of stator-complex interactions with the enteric motor. In the Salmonella enteric motor, the periplasmic region of MotB in a stator complex contacts the P-ring to anchor to the flagellar structure and contacts the peptidoglycan to tether the motor to the cell wall (13). The Vibrio stator complex also interacts with peptidoglycan, but additional interactions with the T-ring allow the stators to be incorporated at a wider position in the Vibrio flagellar motor than in Salmonella. Currently, it is unknown if the C. jejuni stator complexes also interact with peptidoglycan, although the core OmpA peptidoglycan-binding domain motif is intact (47), and MotB from closely related H. pylori has been cocrystallized with a bound glycan strand (48). Our work suggests that MotB also must interact with PflB, because these two proteins colocalize in subtomogram averages, and deletion of pflB results in the failure of MotB to incorporate into the motor, further supporting our conclusion that the disk complex functions to scaffold stator complexes. We did not observe stator-complex densities in the Salmonella motor despite evidence that Salmonella minicells swim (31); this finding is consistent with previous studies that failed to visualize Salmonella stator complexes (18, 31). Because Salmonella stator complexes are dynamic (49), we suspect that under our low-load conditions few are engaged and thus will be difficult to resolve without increasing stator-complex occupancy, using classification approaches, or using larger datasets. Comparing the Salmonella motor with V. fischeri and C. jejuni further suggests that Salmonella incorporates no more than 11 stator complexes, because a stator ring immediately around the rod would be too narrow to incorporate more. This limitation has been noted previously in a model of stator-complex incorporation into the V. alginolyticus motor when the additional stator radius provided by the scaffolding role of the T-ring is not taken into account (16). A Structural Rationale That Quantitatively Accounts for Higher Torque Generation. Considering this work and interpreting previous work (18, 19), we see a compelling correlation between the torque of a flagellar motor and its architecture. In addition to the results reported here, the spirochete flagellar motor with a torque of ∼4,000 pN⋅nm has 16 proposed stator-complex densities (17, 19) in a 30-nm radius ring (although it is curious that the spirochetes have convergently evolved an alternative stator scaffold, the P-collar, which shares no homologous components with the Vibrio or ε-proteobacterial disk complexes). Can these data quantitatively predict observed motor torques? Given that the torque of a single stator complex in enteric bacteria is 146 pN⋅nm (14), our data, consistent with others, show that the lever contact point at which MotA contacts the outer lobe of FliG to apply force is ∼20 nm from the motor axis of rotation (31, 33), leading to the estimation of the force exerted by a single stator complex as ∼7.3 pN. There is good evidence for the assumption that stator complexes exert a constant force in all bacteria, because proton motive forces across bacteria are consistently reported to be between −100 and −200 mV (50); however it should be noted that the sodium motive force in sodium-driven motors may be higher than the proton motive force. Although sodium motive force increases with increasing external sodium ion concentration, it plateaus at an external sodium concentration of ∼250 mM, and the a maximum sodium motive force does not exceed −200 mV (25, 51). Thus, given the radius of stator complexes around the axis of rotation and the number of stator complexes, we can predict the torque of a motor by making the approximation that the torque from multiple stator complexes is roughly additive (14). Our structural data are sufficient to make this prediction for four groups of organisms (Fig. 6A): enteric bacteria such as Salmonella (in which 11 stator complexes exert force at a radius of 20 nm); Vibrio spp. (in which 13 stator complexes exert force at a lever contact point of 21.5 nm); ε-proteobacteria (in which 17 stator complexes exert force at a lever contact point of 26.5 nm); and spirochetes (in which 16 stator complexes exert force at a lever contact point of 30.5 nm). Multiplying the number of stator complexes by the radius of the contact point lever and by the estimated 7.3-pN force exerted per stator complex, we observe excellent correlation between our predicted and the observed torque magnitudes in all four groups (Fig. 6B): a predicted torque of 1,606 pN⋅nm for enteric bacteria vs. an observed torque of 1,260 pN⋅nm (14), with some estimates of 2,000 pN⋅nm (20); a predicted torque of 2,040 pN⋅nm for Vibrio vs. an observed torque of 2,200 pN⋅nm at low sodium concentrations, increasing with higher concentrations; a predicted torque of 3,288 pN⋅nm for ε-proteobacteria vs. an observed torque of 3,600 pN⋅nm; and a predicted torque of 3,562 pN⋅nm for spirochetes vs. an observed torque of 4,000 pN⋅nm. Although this model is clearly a simplification of the process, and biophysical studies reveal that additional stator complexes provide incrementally smaller torque contributions (14), we believe the salient features of our model's predictions are compelling. Another possibility is that ε-proteobacteria assemble more stator complexes at wider radii to accommodate a lower proton motive force. However, the proton motive force of H. pylori has been reported to be relatively high, −220 mV (52). Alternatively, additional stator complexes may act as more sensitive mechanosensors. Fig. 6. Wider stators featuring more stator complexes quantitatively account for torque diversity. (A) Disk complexes scaffold stators to increase the number of stators and the radius of the stator-complex ring. The stator number and increasing wider C-ring in each flagellar motor correlate directly with the torque produced by each motor. (B) Comparison of predicted and observed torques of flagellar motors from various species. Filled circles represent enteric bacteria; open circles represent Vibrio spp.; open squares represent ε-proteobacteria; and filled squares represent spirochetes. Where there was no direct torque measurement, C. jejuni and B. burgdorferi torques are inferred from closely related species with similar motors (H. pylori and Leptospira, respectively). Relative torque strengths are validated by swimming speeds and the ability of different bacteria to swim through viscous fluids (1, 4, 7). The dotted line represents a perfect prediction. In all organisms the C-ring radius tends to scale with both stator-ring radius (as would be expected to maintain MotA:FliG C interaction) and motor torque. Curiously, however, although this correlation is clear, it is not tightly constrained. For example, ε-proteobacteria and spirochetes have wide stator- and C-rings, but the C-ring is wider in spirochetes than in ε-proteobacteria; conversely the stator ring is wider in ε-proteobacterial than in spirochetes. The observation that these variables scale together, but without strict correlation, provides a mechanistic insight: No strict stoichiometric or symmetric correspondence is necessary for motor function. This lack of strict correlation also suggests a clear reducibly complex pathway for the evolution of higher-torque motors, because the increases in C-ring and stator-ring radii can be staggered progressively and asynchronously over evolutionary time through the addition of spacers while a functional motor is maintained. Together these results indicate that different bacteria have modified their motors to produce torques suited to their environments. C. crescentus has evolved a low-torque efficient motor suited to its low-nutrient, low-viscosity freshwater habitat, whereas the ε-proteobacteria and spirochetes have evolved high-torque motors that require high energy expenditure to bore through viscous mucus and tissue. Such modification of the mechanical output of a molecular machine is not without precedent, because similar observations have been made in the unrelated F-ATPase rotor ring (53). Our observations propose a mechanistic model for how different bacterial flagellar motors produce different torques. By assembling wider stator rings using a proteinaceous scaffold, individual stator complexes can exert greater leverage on the axial driveshaft of the motor, producing greater torque. Additionally, more stator complexes can be accommodated around the motor, further increasing torque. In addition to describing the evolution of higher torque, this study illustrates a mechanism for adapting mechanical output that might be capitalized on in future synthetic repurposing of molecular machinery.

Materials and Methods Construction of C. jejuni Mutants. All strains are listed in Table S1, and plasmids used are listed in Table S2. C. jejuni mutants were constructed by electroporation as previously described (54). For removal from the C. jejuni 81-176 SmR (DRH212) chromosome, genes were first amplified by PCR using primers containing 5′ BamHI restriction sites. Each amplified fragment contained the gene of interest with ∼750 flanking regions. Cloning of fragments into pUC19 resulted in plasmids pLKB652 (pUC19::pflB), pDRH2526 (pUC19::pflA), and pDRH2312 (pUC19::motAB). An SmaI-digested cat-rpsL cassette was ligated into the SwaI site of pflB in pLKB652 to create pLKB658, into the EcoRV site of pflA in pDRH2526 to create pDRH2532, and into the SpeI site of motA in pDRH2312 to create pDRH3330. To insert the cat-rpsL cassette in motB, site-directed mutagenesis was used to create an EcoRV site within motB to form pDRH2324 followed by the insertion of cat-rspL to generate pDRH3331. Table S1. Strains used in the study Table S2. Plasmids used in this study Plasmids were electroporated into C. jejuni 81-176 SmR (DRH212) to inactivate each respective gene on the chromosome by insertion of the cat-rpsL cassette. In addition, the previously constructed plasmids pDRH2534, pDRH2536, and pALU101 were used to interrupt flgF, flgI, and flgH, respectively, with a cat-rpsL cassette on the C. jejuni 81-176 SmR (DRH212) chromosome. Transformants were recovered on Mueller Hinton (MH) (Difco) agar containing chloramphenicol and mutants were verified by colony PCR to recover the following isogenic mutants: ABT337 (pflB::cat-rpsL), ALU103 (flgH::cat-rpsL), DAR866 (pflA::cat-rpsL), DRH2550 (flgF::cat-rpsL), DRH2553 (flgI::cat-rpsL), MB1225 (motA::cat-rpsL), and MB1226 (motB::cat-rpsL). In-frame deletions of genes were created with specific primers using PCR-mediated mutagenesis or gene splicing by overlap extension mutagenesis (55, 56). Resulting plasmids were verified by sequencing to form pABT324 (pUC19::ΔpflB), pDRH2745 (pUC19::ΔpflA), pDAR1012 (pUC19::ΔmotA), and pDAR1013 (pUC19:: ΔmotB). Gene deletions were achieved by electroporating corresponding plasmids into strains containing genes interrupted by the insertion of the cat-rpsL cassette. Transformants were recovered on MH agar containing 0.5, 1, or 2 mg/mL streptomycin and then were screened for chloramphenicol sensitivity. Deletions were verified by PCR, which resulted in the creation of isogenic 81-176 SmR mutants containing deletions: ΔpflB (DAR981), ΔpflA (DAR1124), ΔmotA (DAR1131), ΔmotB (DAR1066), ΔflgH (DRH2449), and ΔflgF (SNJ922). Bacterial Growth. Salmonella strain TH16943 (ParaftsZ) was grown aerobically overnight in LB broth from freezer stocks at 37 °C. After overnight culture, 50 μL of 5% (wt/vol) l-arabinose were added to 5 mL fresh LB broth for each culture, and cultures were incubated for an additional 5 h. To enrich for minicells, 3 mL of this culture were spun at 6,000 × g for 5 min, and the supernatant was recovered. The supernatant was spin-concentrated by centrifugation at 18,000 × g for 5 min, and the resultant pellet was resuspended in ∼50 μL LB broth and was immediately plunge-frozen. V. fischeri was grown aerobically overnight at 28 °C in salt-supplemented LB medium (38) (LBS) with 35 mM MgSO 4 . Cultures were reinoculated into fresh LBS, harvested in early log phase, spin-concentrated to an OD 600 of ∼20, and immediately plunge-frozen. C. jejuni were grown for 48–60 h on MH agar under microaerobic conditions using CampyPak sachets (Oxoid) at 37 °C. Cultures were generously restreaked and returned to incubate for a further 16 h. Bacteria were harvested by resuspension into ∼1 mL MH broth to an OD 600 of ∼10.0 and were immediately plunge-frozen. Preparation of Electron Microscopy Samples, Data Collection, and Tomogram Reconstruction. Quantifoil R2/2 grids (200-mesh) (Quantifoil Micro Tools GmbH) were glow-discharged for 60 s at 10 mA. A solution of 10-nm colloidal gold was added to cells immediately before plunge freezing. A 3-μL droplet of the sample solution was applied to the glow-discharged electron microscopy grid; then the grid was blotted and plunge-frozen into a liquid ethane-propane mixture using a Vitrobot plunge-freezing robot (FEI Company) using a wait time of 60–180 s, a blot time of 5–10 s, and blot offsets between −2 and −5 mm. Grids were stored under liquid nitrogen until data collection. Tilt series were collected on a 200-kV FEI Tecnai TF20 FEG transmission electron microscope (FEI Company) equipped with a Falcon II direct electron detector camera (FEI Company) using Gatan 914 or 626 cryo-holders. Tilt series were recorded from −61° to +61° with an increment of 3° collected defocus between −4 μm and −10 μm using Leginon automated data-collection software (57) at a nominal magnification of 50,000× and were binned four times to final pixel size of 0.81nm. Cumulative doses of ∼120 e−/Å2 were used. Overnight data collection was facilitated by the addition of a 3-L cold-trap Dewar flask and automated refilling of the Dewar cryo-holder triggered by a custom-written Leginon node interfaced with a computer-controlled liquid nitrogen pump (Norhof LN2 Systems). Tomograms were reconstructed automatically using RAPTOR (58) and the IMOD package (59). Low-defocus images were low-pass filtered to remove data beyond 3.5 nm−1. High-defocus datasets were contrast transfer function (CTF)-corrected using TomoCTF (60). Subtomogram Extraction, Alignment, and Averaging. Positions of flagellar motors in tomograms were initially aligned manually along their rotational axes. Datasets were halved, and the particle estimation for electron tomography (PEET) package (61) was used for iterative subtomogram extraction, fine alignment, and averaging of the two halves independently. Resolution was estimated by gold-standard Fourier shell correlation (FSC) by correlating the two halves of the dataset using FSC (Fig. S1). Salmonella motors and C. jejuni ΔpflA, ΔpflB, ΔflgP, and ΔflgQ exhibited flexibility between the top and bottom parts of the motor. In these cases individual averages were constructed for the top and bottom segments and ultimately were merged into a composite structure in analogy to recent single-particle studies (62). Tn Mutagenesis and Screening of C. jejuni 81-176. To identify potential disk mutants of C. jejuni 81-176, we conducted a three-step Tn mutagenesis screening procedure to identify flagellated but nonmotile mutants. C. jejuni 81-176 SmR ΔastA flaB::astA chromosomal DNA was used in in vitro transposition reactions with the darkhelmet Tn as described previously (54, 63, 64). Tn mutants were recovered on MH agar containing chloramphenicol, kanamycin, and 35 μg/mL 5-bromo-4-chloro-3-indolyl sulfate. Approximately 8,000 colonies displaying a blue phenotype indicating an intact transcriptional pathway for the expression of flagellar genes were stabbed in 0.4% MH motility agar and incubated for 24 h in microaerobic conditions. Mutants with impaired motility were recovered and grown for an additional 48 h. Mutants then were restreaked, grown for another 16 h, and resuspended to an OD 600 of 0.8 in MH broth in 15-mL conical tubes. After at least 1-h incubation at room temperature, mutants that aggregated (indicating production of flagella), as determined by visual inspection of tubes, were saved to identify the location of the Tn insertion by direct DNA sequencing with a primer annealing to the end of the Tn. Antisera Generation. Primers were designed to amplify the coding sequence of motA from codon 2 through the stop codon with in-frame BamHI and XhoI restriction sites added to the 5′ ends of the primers. PCR-amplified motA was ligated into pET21a digested with BamHI and XhoI, which allowed expression of MotA with a C-terminal 6×His -tag in pDAR1180. Primers were designed to amplify the coding sequence of motB from codon 2 through the penultimate codon with in-frame NdeI and PstI restriction sites added to the 5′ end of the primers. In addition, one primer contained codons for a C-terminal 6×His -tag to be added to the penultimate codon on motB. PCR-amplified motB was ligated into pT7-7 digested with NdeI and PstI, which allowed the expression of MotB-6×His and created pDAR906. Primers were designed to amplify the coding sequence of pflB from codons for V103 to L304 with in-frame BamHI and PstI sites added to the 5′ end of the primers. Additionally, one primer contained codons for a C-terminal 6×His tag to be added after codon 304. After digestion, the PCR-amplified pflB fragment was ligated into BamHI- and PstI-digested pMAL-c2X, allowing expression of a fragment of PflB V103-L304 fused to maltose-binding protein (MBP) with a C-terminal 6×His tag to create pDAR2226. Primers were designed to amplify the coding sequence of pflA from codon 2 to the stop codon with in-frame BamHI and PstI sites added to the 5′ end of the primers. In addition one primer also contained codons for a C-terminal 6×His tag to be added to the last codon. After digestion, PCR-amplified pflA was ligated into BamHI- and PstI-digested pMal-p2X, which allowed expression of PflA fused to MBP with a C-terminal 6×His tag to create pDAR2111. For the induction of MotA-6×His, MotB-6×His, PflB V103-L304 -MBP-6×His, and PflA-MBP-6×His, the respective plasmids were transformed into BL21 (DE3). Bacteria were grown in 500 mL LB to an OD 600 of 0.4. For induction, isopropyl β-d-1-thiogalactopyranoside was added to a final concentration of 1 mM, and the cultures were incubated for an additional 3 h. Bacteria expressing recombinant proteins were harvested by centrifugation and lysed with an EmulsiFlex C5 cell disrupter (Avestin). Lysis buffer for purification of MotB-6×His contained 0.5 M sucrose and 0.2% Zwittergent (Calbiochem). Lysis buffer for MotA-6×His purification contained 0.5 M sucrose, 0.2% Zwittergent, and 8 M urea. PflA-MBP-6×His and PflB V103-L304 -MBP-6×His proteins were purified in denaturing conditions with 8 M urea. All proteins were purified by Ni-NTA chromatography. Purified proteins were used to immunize mice or guinea pigs for the generation of polyclonal antisera. Fractionation of C. jejuni Strains and Analysis of Proteins. Recovery of whole-cell lysates and fraction of C. jejuni strains into subcellular compartments for analysis of localization of proteins from the inner membrane, outer membrane, periplasm, and cytoplasm were performed as previously described (65). Proteins from each fraction representing equivalent cell numbers (∼200 μL of a C. jejuni culture at an OD 600 0.8) were loaded onto 10% SDS/PAGE gels and then were transferred to membranes for immunoblot analysis. For immunoblotting analyses, the following primary antisera were used at the stated dilutions: FlgP M4 or M5 (1:2,000) (41), MotA M219 (1:500), MotB M195 (1:500), PflA M231 (1:500), and PflB GP143 (1:500). Secondary antisera were used at a dilution of 1:15,000.

Acknowledgments We thank Anchi Cheng for advice on programming an additional Leginon node; Kelly Hughes for the generous gift of the Salmonella minicell strain TH16943; Tillmann Pape and Amanda Wilson for technical assistance during electron microscopy data collection; and Bonnie Chaban, Velocity Hughes, Ariane Briegel, Alain Filloux, and Richard Berry for critical reading of the manuscript. This work was supported by Biotechnology and Biological Sciences Research Council Grant BB/L023091/1 (to M.B.), Howard Hughes Medical Institute funding (G.J.J.), and National Institutes of Health Grants 5R01AI065539 and 5R21AI103643 (to D.R.H.).

Footnotes Author contributions: M.B., D.A.R., G.J.J., and D.R.H. designed research; M.B. and D.A.R. performed research; M.B., C.A.B., and E.G.R. contributed new reagents/analytic tools; M.B., G.J.J., and D.R.H. analyzed data; and M.B., G.J.J., and D.R.H. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Data deposition: The electron cryo-tomography subtomogram average density maps reported in this paper have been deposited in the Electron Microscopy Data Bank (EMD) (accession nos. Salmonella WT: EMD-3154; Vibrio fischeri WT: EMD-3155; Campylobacter jejuni WT: EMD-3150; V. fischeri motB: EMD-3156; C. jejuni motB: EMD-3157; C. jejuni flgQ: EMD-3158; C. jejuni flgP: EMD-3159; C. jejuni pflA: EMD-3160; C. jejuni pflB: EMD-3161; and V. fischeri flgP: EMD-3162).

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