The V1- and F1- rotary ATPases contain a rotor that rotates against a catalytic A3B3 or α3β3 stator. The rotor F1-γ or V1-DF is composed of both anti-parallel coiled coil and globular-loop parts. The bacterial flagellar type III export apparatus contains a V1/F1-like ATPase ring structure composed of FliI6 homo-hexamer and FliJ which adopts an anti-parallel coiled coil structure without the globular-loop part. Here we report that FliJ of Salmonella enterica serovar Typhimurium shows a rotor like function in Thermus thermophilus A3B3 based on both biochemical and structural analysis. Single molecular analysis indicates that an anti-parallel coiled-coil structure protein (FliJ structure protein) functions as a rotor in A3B3. A rotary ATPase possessing an F1-γ-like protein generated by fusion of the D and F subunits of V1 rotates, suggesting F1-γ could be the result of a fusion of the genes encoding two separate rotor subunits. Together with sequence comparison among the globular part proteins, the data strongly suggest that the rotor domains of the rotary ATPases and the flagellar export apparatus share a common evolutionary origin.
Citation: Kishikawa J-i, Ibuki T, Nakamura S, Nakanishi A, Minamino T, Miyata T, et al. (2013) Common Evolutionary Origin for the Rotor Domain of Rotary Atpases and Flagellar Protein Export Apparatus. PLoS ONE 8(5): e64695. https://doi.org/10.1371/journal.pone.0064695
Editor: Hendrik W. van Veen, University of Cambridge, United Kingdom
Received: February 13, 2013; Accepted: April 17, 2013; Published: May 28, 2013
Copyright: © 2013 Kishikawa et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was partly supported by Grants-in-Aid from the Ministry of Education, Science, Sports and Culture of Japan (No. 21370042 and 24370059 to KY, and 18074006 to KI), Targeted Proteins Research Program (TPRP) (B-37, to KY and FBA1 to KI). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Two types of rotary ATPases are found in biological membranes; VoV1(V type ATPase) and FoF1 (F type ATPase) –. Evolutionary counterparts of the eukaryotic VoV1 are found in most archea and some bacteria (often referred to as A-type ATPase). Both VoV1 and FoF1 couple ATP synthesis and hydrolysis to proton translocation across the membrane by rotation of the rotor apparatus against the surrounding stator which includes the catalytic A3B3 or α3β3 hexamer (Fig. 1a). Sequence and structural comparison of the two ATPases indicate significant homology between the catalytic subunits, but not between the subunits of the central rotor domain . For example, VoV1 lacks a counterpart of the rotor shaft F1-γ subunit , , .
(a) Schematic model of prokaryotic FoF1 (left) and VoV1 (right). Rotor subunits (D, F, Vo-d in VoV1, γ and ε in FoF1) are presented in brown. Rotor rings composed of hydrophobic c subunit are presented in dark brown. Peripheral stators (EG and Vo-a in VoV1, b2, δ, and Fo-a in FoF1) are presented in sky blue. Schematic model of the flagellar type III export apparatus (lower). The export apparatus consists of a proton-driven export gate made of six integral membrane proteins, FlhA, FlhB, FliO, FliP, FliQ and FliR, and a water-soluble ATPase complex composed of FliH, FliI, and FliJ. The FliI6-FliJ shows remarkable structural similarity to F1 and V1. The amino acid sequence of FliH shows sequence similarity to peripheral stalk E subunit of VoV1. (b) Crystal structure of γ subunit from bovine FoF1 (PDB: 1E79). (c) Crystal structure of DF subunit of E.hirae VoV1 (PDB: 3AON). (d) Crystal structure of FliJ of Salmonella typhimurium (PDB:3AJW).
The minimal ATP-driven rotary unit of FoF1 is F1, which is comprised of three different proteins with a stoichiometry of α3β3γ. The F1-γ contains two distinct domains; a coiled-coil domain that penetrates the central cavity of the α3β3 cylinder from the top to the bottom, and a globular domain containing a α/β fold which makes contact with the F1-ε (Figs. 1a,b).
For VoV1, the minimal rotary unit consists of the A3B3D subunits . In addition, analysis of the secondary structure of the D subunit predicts the presence of long α-helices at both the amino and carboxyl termini as found in the F1-γ (Fig. 1b, Supplementary Fig. S1a). On the basis of this, it has been suggested the D subunit is likely to be a structural and mechanistic analog of F1-γ despite the lack of any significant sequence similarity . However, FoF1 lacks a counterpart of the V1-F subunit , , , which has a typical globular α/β fold  (Fig. 1c). Intriguingly, the X-ray structure of the entire V1 of T. thermophilus revealed that the central rotor subunit contains both the coiled-coil (D) and globular domains (F), suggesting that the V1-F and V1-D subunits together form the counterpart of the α/β domain of F1-γ .
Mulkidjanian et al. have proposed a scenario to explain the origin of rotary ATPases, where rotary ATPases share an evolutionary origin with the bacterial flagellar and non-flagellar type III export systems . The flagellar export apparatus consists of a membrane-embedded export gate composed of FlhA, FlhB, FliO, FliP, FliQ and FliR, and a water-soluble ATPase complex consisting of FliH, FliI, and FliJ (Fig. 1a, lower panel) . Components of the flagellar ATPase complex, which allows the export gate to efficiently utilize proton motive force across the cytoplasmic membrane as an energy souse for protein translocation , , exhibit extensive structural and sequence similarity to catalytic and rotor subunits of rotary ATPases. For instance, the atomic structure of FliI ATPase of the flagellar export apparatus is remarkably similar to the F1 β/α and V1 A/B subunits . FliJ, which is a soluble export component protein, also shows a striking structural similarity to the coiled-coil region of F1-γ . This structural motif is also seen in the YscO-like protein, CT670 from Chlamydia tracomatis, a FliJ homolog of the non-flagellar type III export apparatus . Interestingly, FliJ promotes the formation of a homo-hexamer of FliI by binding to the center of the ring, facilitating the FliI ATPase activity . A very similar arrangement is observed in the FoF1-ATPase with the antiparallel α-helical coiled coil formed by the N- and C-terminal regions of the γ subunit penetrating into the central cavity of the α3β3 ring. These findings indicate that the type III export system has a F1- or V1-like structure and share a common evolutionary origin.
Here we aimed to explore the evolutionary relationship between the rotor domains of the VoV1 and FoF1 and the type III export system. FliJ formed chimeric complex with A3B3 of VoV1 and promoted their ATPase activity. Single molecular analysis indicates anti-parallel coiled coil structure (FliJ structure) protein functions as a rotor axis. These results strongly suggest that the FliJ structure proteins are the ancestral subunit of the rotor subunit of rotary ATPases. We also discuss the evolutionary relationship between globular domain of F1-γ, and V1-F.
The A3B3JL and A3B3DFf expression plasmids were constructed from His-tagged V1 (A(His-10/C28S/S232A/T235S/C255A/C508A)3B(C264S)3D(E48C/Q55C)F) as described in Figs. S3a, S4a. The A3B3JL (A(His-10/C28S/S232A/T235S/C255A/C508A)3B(C264S)3JL), A3B3DFf, and A3B3 were expressed in E. coli and the expressed enzymes purified by Ni2+-affinity chromatography (Qiagen) followed by ion exchange on a RESOURCE Q column (GE healthcare) . The purified His-tagged enzymes were biotinylated at two cysteines using 6-[N-[2-(N-maleimide)ethyl]-N-piperazinylamide]hexyl-D-biotinamide (Dojindo, Kumamoto, Japan). The bound ADP in each enzyme was partially removed by successive EDTA-heat treatment .
FliJ was expressed in BL21(DE3)pLysS and purified as described previously . FliJ(C32T/I61C/I67C) was created by QuikChange mutagenesis (Stratagene), expressed in BL21(DE3)pLysS, biotinylated and purified as the same protocol used for FliJ(C32T/I67C) . A3B3 was expressed and purified as previously described . The A3B3-FliJ complex was reconstructed from the purified FliJ and A3B3. FliJ and A3B3 were mixed and incubated overnight at 23°C. The mixture was applied to a HilLoad Superdex 200 column in 20 mM MOPS-NaOH pH 7.0 and 100 mM NaCl. The fraction containing the A3B3-FliJ complex was pooled and stored for future use.
Biotinylated FliJ(C32T/I61C/I67C) and His-tagged A3B3 were mixed and incubated overnight at 23°C. The A3B3-FliJ mixture was applied to 5-ml polypropylene column containing Ni-NTA agarose (Qiagen), and A3B3-FliJ complex was eluted with a sodium phosphate buffer (pH 7.2) containing 200 mM imidazole and 300 mM NaCl. BSA buffer, which is buffer A (50 mM Tris-Cl, pH 8.0, 100 mM KCl, 2 mM MgCl2) containing 2 mg ml−1 BSA, was infused into the flow cell to prevent nonspecific binding, and then the A3B3-FliJ complexes diluted 1∶10 into BSA buffer were attached on the Ni-NTA glass and incubated for 15 minutes at 23°C. Unbound molecules were washed out with BSA buffer, and streptavidin-coated gold spheres with a diameter of 40-nm (BioAssay Works) diluted into BSA buffer and infused into the flow cell. After incubation for 15 minutes at 23°C, unbound gold spheres were washed out with buffer A. After infusion of buffer A containing 4 mM Mg-ATP, 2.5 mM phosphoenol pyruvate, and 0.5 mg ml−1 pyruvate kinase, rotation of the gold spheres were observed by a dark-field microscope (BX53, U-DCW, UPlanFLN 100×, PE 5×; Olympus) and recorded with a high-speed camera (ICL-B0620M-KC, IMPERX) at 0.8–1.6-ms intervals at 23°C. For the rotation assay of A3B3DFf or A3B3JL, the biotinylated enzyme (1–5 nM) in buffer A was applied to the flow cell and incubated for a few minutes at 23°C. Streptavidin-coated magnetic beads (100–300 nm) and Ni2+-NTA coated cover glasses were prepared as previously described , . Unbound enzyme was washed out with 20 µl of buffer A. Then, 20 µl of buffer A with 2 mg ml−1 BSA was infused to the flow cell and incubated for <30 s to prevent nonspecific binding. The BSA solution in the chamber was washed out with 20 µl of buffer A. Then, buffer A containing streptavidin coated magnetic beads (1010<1011 particles ml−1) were infused into the flow cell and incubated for a few minutes. Unbound beads were washed out with 20 µl of buffer A. After infusion of 80 µl of buffer A containing Mg-ATP at the indicated concentration, 2 mM MgCl2, 2.5 mM phosphoenol pyruvate, and 0.5 mg ml−1 pyruvate kinase, rotation of the bead was recorded with a high speed camera (Eclips, IN) at 1000 frames per second (f.p.s.) using a phase-contrast microscope (IX70, Olympus) with ×100 objective lens (N.A., 1.30, Olympus) at 23°C. Images were captured as an 8-bit AVI file. The centroid of the bead images was calculated , .
Electron cryo-microscopy and image analysis
Sample grids were prepared by applying 3 µl of a protein solution containing the in vitro reconstructed A3B3J rings (150 µg/ml) onto a holey carbon grid (Quantifoil R0.6/1.3, Quantifoil Micro Tools, Jena, Germany), which had been glow discharge for 20 s before use. The grids were blotted onto filter paper for 5 s to remove excess solution, vitrified in liquid ethane at −196°C using a Vitrobot (FEI, Eindhoven, Netherlands) and transferred into a liquid nitrogen storage capsule. Particle images were recorded with a 4 K×4 K Slow Scan CCD camera (TemCam-F415MP, TVIPS) mounted on a JEM-3200FSC electron microscope (JEOL, Tokyo, Japan), equipped with a liquid-helium cooled specimen stage, a Ω-type energy filter and a field-emission electron gun operated at an accelerating voltage of 200 kV. Electron micrographs were collected at 50 K with a magnification of ×140,000, corresponding to 1.07 Å/pixel. Focal pairs of the micrographs were recorded at a defocus between 1.5 and 2.5 µm for the first micrograph and at a defocus between 3.5 and 5.5 µm for the second. Electron dose was set to 30 e−/Å2 for both micrographs. The particle images were processed using the EMAN software package . The focal pairs were merged using FOCALPAIR , . The defocus amplitude, envelope and noise values were determined using CTFIT . Micrographs showing significant astigmatism or drift were discarded. The particle images were selected with BOXER  and the boxed particle images were aligned and classified using REFINE2D.PY .
Phylogenetic tree analysis
All sequences used in this study were aligned using the MAFFT program (http://mafft.cbrc.jp/alignment/server/) . Phylogenetic and molecular evolutionary analyses were conducted using MEGA version 5 . The phylogenetic tree was constructed using the Maximum-Likelihood (ML) method under the Jones-Taylor-Thornton model by using MEGA.
Reconstitution of A3B3J
Structural sequence alignment has previously revealed that FliJ shares some amino acid conservation with F1-γ . The structural and sequence similarity suggests that FliJ may behave as a counterpart of rotor subunit of rotary ATPases. To test this possibility, we investigated whether FliJ binds to the center of the A3B3 ring of T. thermophilus, forming the A3B3J complex. A3B3 was mixed with excess amounts of FliJ, and the mixture incubated overnight at room temperature. The mixture was applied to a gel filtration column to remove free FliJ. The retention time of the complex peak was almost identical to that of A3B3 alone (Fig. S2a). SDS-PAGE analysis revealed that FliJ co-eluted with A3B3, indicating the formation of the A3B3J complex (Fig. 2a). Previous studies have shown that V1 and A3B3D are resistant to ATP-induced dissociation  while A3B3 completely dissociates into monomeric A and B subunits by incubation with 1 mM ATP for 30 min (Fig. 2b). In contrast, the A3B3J complex is at least partially resistant to ATP-induced dissociation (Fig. 2b), indicating that the presence of FliJ stabilizes the A3B3 complex to a significant degree. ATP hydrolysis by V1 or A3B3D proceeds at a steady rate for a few minutes, then decelerates slowly, due to ADP inhibition . In contrast, the ATPase activity of A3B3 decelerates rapidly, reaching a steady state rate within a few minutes owing to rapid ATP-induced dissociation of A3B3 . In the case of A3B3J, the ATP hydrolysis profile is similar to that of V1 or A3B3D exhibiting continuous ATP hydrolysis activity after an initial burst phase (Fig. 2c). The turnover rate of A3B3J is ∼7.0 s−1, almost twice that of A3B3 (∼4.0 s−1). Together, these results suggest that FliJ stabilizes the A3B3 hexamer by binding to A3B3 and promotes continuous ATPase activity in a similar manner to the V1-D subunit.
(a) SDS-PAGE analysis of A3B3J and A3B3DFf. (b) Analysis of disassembly of A3B3 and A3B3J on Native PAGE. Each complex was incubated without nucleotide, or with 1 mM MgATP at 37°C for 1 h, followed by separation by Native PAGE. (c) ATP hydrolysis activity of A3B3 and A3B3J. Time courses of ATP hydrolysis catalyzed by A3B3 (blue lines) and A3B3J (red lines) at 25°C and at 4 mM MgATP. The reaction was started by the addition of 20 µl of 1 µM enzyme solution to 2 ml of assay mixture.
Electron cryo-microscopy analysis of A3B3J
In order to identify the location of FliJ in the A3B3J complex, we analyzed the images of frozen-hydrated A3B3J and A3B3 particles embedded in vitreous ice by electron cryo-microscopy. Since the distribution of the particle orientation was strongly biased to end-on view, we aligned and averaged the end-on images of each particle. The averaged image of A3B3 shows a hetero-hexameric ring structure with an unoccupied central hole of 2 nm in diameter (Fig. 3a), consistent with the crystal structure of A3B3 . In contrast, the averaged image of A3B3J, clearly shows extra density in the interior of the central hole but located off center, close to one of the peripheral subunits (Fig. 3b). These observations strongly suggest that FliJ penetrates into the central hole of the A3B3 complex in a way similar to the D-subunit of V1-ATPase .
Rotary motion of FliJ in A3B3 without uni-directionality
We attempted to demonstrate the rotary motion of FliJ against A3B3 immobilizing A3B3J on a Ni-NTA-coated glass surface through His10-tags introduced at the N terminus of the A subunits, and attaching a 40-nm streptavidin-coated gold colloid (40-nm bead) to the biotin-labelled FliJ (Fig. 4a). Beads were imaged by dark-field microscopy, and beads motions were recorded on a fast-framing CCD camera at speeds up to 1,000 frames s−1. We found rotating beads attached to the FliJ in A3B3. It was confirmed that gold beads observed under the microscope were attached to the A3B3J complex through FliJ because very few beads were found without A3B3J. Two to ten beads showing rotary motion were usually found in a single flow cell. However, they did not show apparent uni-directionality. For instance, both rotating beads showing clockwise or anti clockwise rotation (Fig. 4 b–e) were found. Some beads showed stepwise rotary motion (Fig. 4 f,g). The rotary motion of A3B3J was also observed without ATP in the infusion buffer but the number of beads showing rotary motion apparently decreased; one to three beads showing rotary motion were found in a single flow cell. These results indicate that FliJ in the A3B3 does not function as a perfect rotor axis. FliJ is only weakly bound to A3B3, thereby dissociating FliJ from the complex after successive gel filtration chromatography of the A3B3J complex (Fig. S2b). In agreement with this, the ATPase activity of A3B3J (7.0 s−1), although significantly higher than A3B3, is much lower than that of V1 (65 s−1) or A3B3D (25 s−1), reflecting the relative instability of the A3B3J complex. Therefore, the low binding affinity of the FliJ for A3B3 is likely to reduce the chance of FliJ rotation being coupled with a conformational change of A3B3 by ATP hydrolysis.
(a) Schematic representation of the experimental system. A 40 nm gold particle was attached to FliJ via a streptavidin-biotin linkage. Time courses of the anti-clockwise (b, c) and clockwise rotation (d, e) of beads attached to the shaft of A3B3J captured at speeds up to 1,000 frames s−1 at 2 mM [ATP]. Stepping motions of beads were shown in f and g. Trajectories of the bead centroid are shown in insets.
Anti-parallel coiled coil structure (FliJ structure) functions as a rotor
FliJ adopts an anti-parallel coiled coil structure composed of both N- and C-terminal α-helices and is similar to that of V1-D or F1-γ . If we assume that the rotor subunits of the rotary ATPases and the flagellar type III export apparatus share a common evolutionary origin, then the ancestral rotor subunits adopt an anti-parallel coiled-coil structure, that is, the FliJ structure. In order to demonstrate that the FliJ structure functions as a rotor, we constructed a gene encoding a FliJ structure (FliJ-like protein, termed JL hereafter). The JL gene encodes an anti-parallel coiled coil region of V1-D composed of both N- and C-terminal α-helices, without the additional region (52 a.a. length) between the N- and C-terminal helices (Fig. S1a). Then we constructed an expression vector encoding the JL gene along with atpA and atpB genes (Fig. S3a). The JL protein was co-expressed along with the A and B subunits in Escherichia coli and purified the A3B3JL complex to homogeneity (Fig. 2a). The A3B3JL was active as an ATPase and the Vmax value of the A3B3JL is similar to that of A3B3D (Fig. S3b). Using a direct observation system similar to that described for V1 , the rotation of the A3B3JL was able to be visualized (Fig. 5a). We observed rotating beads attached to the A3B3JL when the flow cell was infused with buffer containing 4 mM or 10 µM ATP (hereafter [ATP]) (Fig. 5b). The results clearly indicate that the JL protein functions as a rotor. At 10 µM [ATP], A3B3JL showed a clear stepwise rotation, pausing every 120° like F1 or V1. Interestingly, frequent backward steps were observed in rotation of A3B3JL but were seldom seen in stepwise rotation of either V1 or F1, indicating that the extra 53 amino acid residues inserted between the two helices were necessary for continuous rotation without irregular motions (Fig. 5b, inset). Taken together, we propose that an anti-parallel coiled-coil fold unit like FliJ is sufficient to function as a rotor for the ATP-driven rotary motor.
Schematic representation of the experimental system for rotation of A3B3JL (a) and A3B3DFf (c). A magnetic bead was attached to rotor subunit via a streptavidin-biotin linkage. Typical time courses of the rotation of beads attached to the shaft of A3B3JL (b) and A3B3DFf (d) captured at 1000 frames s−1 at 10 µM (red lines) and 2 mM [ATP] (blue lines). In insets, the most populated angles for each enzyme are shown by the horizontal lines separated by 120°. In b, the movements corresponding to backward step of A3B3JL are indicated by dashed arrows.
F1-γ like rotor derived from V1-D and V1-F
As shown in Figure 1, the similar folds of V1-F and the globular domain of F1-γ prompted us to probe an evolutionary relationship between these two proteins, i.e if could F1-γ be derived from two separate proteins as observed for V1-D and V1-F. To investigate the functional relationship between V1-F and globular domain of F1-γ, we constructed an expression vector of a mutant V1 containing a fusion of the genes coding for subunits D and F. The gene coding for F (atpF) was inserted between the two helical regions of the gene coding for D (atpD; see Fig. S4a) and the resulting fusion protein was termed DFf. The mutant ATPase containing the fused protein (A3B3DFf) was expressed in E. coli and purified to homogeneity (Fig. 2a). The A3B3DFf showed an ATPase activity and exhibited simple Michaelis-Menten kinetics, nearly equal to those of wild type V1, which contains separate D and F subunits in the rotor . Using a direct observation system similar to that described for V1, rotation was visualized via a bead attached obliquely to the helical region in the DFf (Fig. 5c). We observed rotating beads attached to the A3B3DFf at 4 mM or 10 µM ATP (Fig. 5d). Stepwise rotation of the A3B3DFf pausing every 120° was also observed at 10 µM ATP. In this case the frequent backwards steps seen for A3B3JL were not observed (Figs. 5d, inset). Together, the identical rotation behavior and kinetic parameters of A3B3DFf confirm that the DFf fully functions as a shaft in the rotary motor. These observations indicate that the V1-F when fused to V1-D, functions in the rotor in the same way as the globular domain of F1-γ and suggest an evolutionary relationship between V1-F and V1-D and F1-γ. In other words, F1-γ could be the product of fusion of the genes encoding an ancestral helical protein similar to V1-D and an ancestral globular protein similar to V1-F.
In this study, we have provided several lines of evidence on the functional similarity of FliJ of the flagellar type III export system to the rotor subunit in rotary ATPases. Analysis of reconstituted A3B3J revealed that FliJ stabilizes the A3B3 hexamer by penetrating into A3B3 and promotes continuous ATPase activity in a manner similar to the V1-D subunit. Although FliJ in A3B3 does not show unidirectional rotation coupling with ATP hydrolysis due to its low binding affinity for A3B3, The JL protein is sufficient for functioning as a rotor of the ATP-driven rotary motor. Based on very recent high resolution X-ray crystal structure of V1 from E. hirae, Arai and co-workers have proposed that the interaction of the coiled-coil part of V1-D with the A subunit is essential for rotation of V1-D against A3B3 hetero-hexamer . In their V1 structure, the globular-loop part of V1-DF is rarely in contact with the A3B3 hexamer. Our results, in which the anti-parallel coiled-coil structure is sufficient as a rotor in rotary ATPases, are consistent with their structural study.
Here we propose that V1-D, F1-γ and FliJ evolved from a common evolutionary origin. The JL protein derived from V1-D functioned as a rotor shaft (Figure 5A), strongly suggesting that FliJ maintains features of a prototype rotor. Because the rotor is composed of separate helical and globular subunits in VoV1, this rotor is an intermediate between the ancestral rotor and the F1-γ, which is a single protein containing both helical and globular domains. Therefore, we propose a possible scenario of evolutional process of rotor apparatus in rotary motors (Fig. 6). In this scenario, F1-γ evolved from a gene fusion of genes encoding the ancestral V1-D and V1-F like proteins. However, because there is the sequence and structural diversity between V1-F and in the globular domain of F1-γ. we cannot exclude the possibility that V1-F is not ancestral gene of globular domain of F1-γ.
V1-D is evolved from a FliJ like protein by adding loop region. V1-D forms a separate rotor shaft together with the globular V1-F. F1-γ evolved through gene fusion of two genes encoding the ancestral V1-D and globular proteins. Assuming that VoV1 have conserved the ancestral form in the rotor apparatus, C subunit (Vo-d) has been replaced with F1-ε during evolution of FoF1. Function of each rotor subunit was summarized in Table S1.
V1-F is structurally similar to CheY, a regulatory subunit of the bacterial flagellar motor, which functions to switch the direction of rotation . A phylogenetic tree analysis using the Maximum Likehood (ML) method indicated that V1-F is evolutionarily related to CheY rather than to the globular domain of F1-γ (Fig. S5). The topology of V1-F is also more similar to that of CheY. (Fig. S5). This indicates that V1-F and CheY share a common evolutionary origin. In contrast, the sequence and structural diversity between V1-F/CheY and in the globular domain of F1-γ can be explained. The central rotor apparatus of the two rotary ATPases contain significant structural differences. The Vo sector contains the funnel shaped C subunit (Vo-d subunit), which serves as a socket for the DF rotor in V1 , . In contrast, F1-γ attaches directly onto the Fo-c ring while the F1-ε forms contacts with both the F1-γ and Fo-c ring ,  (see Fig. 1a). It is possible that the differences in contact features between V1-F and the globular domain of F1-γ have promoted structural and sequence diversity of the rotors during the evolution of the two different ATPases.
In contrast to V1-DF and the F1-γ, there is little similarity between the other central rotor domain of the VoV1- and FoF1 (see Fig. 1a). The F1-ε, composed of an N-terminal β sandwich and a short C terminal helix ,  shows neither sequence nor structural similarity to the equivalent Vo-d (prokaryotic C subunit). Assuming that VoV1 have conserved the ancestral form in the rotor apparatus, Vo-d has been replaced with F1-ε during evolution of FoF1.
Secondary structure prediction of the D subunit of T. thermophilus V-ATPase (a), the γ subunit of E. coli F1. (b) and FliJ of S. enterica (c) using PORTER: http://distill.ucd.ie/porter/. Predicted helical, sheet, and coiled regions are indicated by H, E, and C, respectively. For the γ subunit, both N- and C-terminal helices in the crystal structure (PDB: 1E79) are indicated by black lines. For the D subunit, assigned helices in the crystal structure (PDB: 3A5C) are indicated by black lines. Other regions are disordered in the crystal structure. For the FliJ, both N- and C-terminal helices in the crystal structure (PDB: 3AJW) are indicated by black lines. The site for insertion of the F subunit in the D subunit is indicated with red characters.
(a) Analysis of ATPase complexes with gel-permeation chromatography. The mixture of A3B3 and FliJ was incubated at room temperature for overnight, and then applied onto Superdex-200 equilibrated with 20 mM MOPS (pH 7.0) and 150 mM NaCl. (b) SDS-PAGE analysis for A3B3J after successive gel-permeation chromatography. A3B3J was further applied onto gel-permeation chromatography, the resultant complex was analyzed by 15% SDS-PAGE (see lane marked 2nd).
(a) Construction of the A3B3JL expression vector. (b) ATP hydrolysis activity of A3B3JL at the indicated [ATP].
(a) Construction of A3B3DFf expression vector. (b) ATP hydrolysis acitivity of A3B3DFf at the indicated [ATP].
Phylogenetic tree of V1-F (red circle), the globular domain of F1-γ (blue circles), and CheY (green circles). Open circles indicate genes from eukaryotes. Construction of the phylogenetic tree is described in the Methods section.
We thank S. Furuike, S. Akimoto, K. Tanata, M. Tanigawara, members of the Yosida, Kinosita, Noji labs and ICORP in Odaiba for help and advice, and Dr. Bernadette Byrne and May K. Macnab for critical reading of the manuscript.
Conceived and designed the experiments: KY KI. Performed the experiments: JK IT AN SN HU HK. Analyzed the data: JK IT AN SN HU HK. Contributed reagents/materials/analysis tools: KN. Wrote the paper: KY KI TM.
- 1. Forgac M (2007) Vacuolar ATPases: rotary proton pumps in physiology and pathophysiology. Nat Rev Mol Cell Biol 8: 917–929.
- 2. Yoshida M, Muneyuki E, Hisabori T (2011) ATP synthase–a marvellous rotary engine of the cell. Nat Rev Mol Cell Biol 2: 669–677.
- 3. Junge W, Sielaff H, Engelbrecht S (2009) Torque generation and elastic power transmission in the rotary F(O)F(1)-ATPase. Nature 459: 364–70.
- 4. Yokoyama K, Imamura H (2005) Rotation, structure, and classification of prokaryotic V-ATPase. J Bioenerg Biomembr 37: 405–410.
- 5. Mulkidjanian AY, Makarova KS, Galperin MY, Koonin EV (2007) Inventing the dynamo machine: the evolution of the F-type and V-type ATPases. Nat Rev Microbiol 5: 892–899.
- 6. Imamura H, Ikeda C, Yoshida M, Yokoyama K (2004) The F subunit of Thermus thermophilus V1-ATPase promotes ATPase activity but is not necessary for rotation. J Biol Chem 279: 18085–18090.
- 7. Makyio H, Iino R, Ikeda C, Imamura H, Tamakoshi M, et al. (2005) Structure of a central stalk subunit F of prokaryotic V-type ATPase/synthase from Thermus thermophilus. EMBO J 24: 3974–3983.
- 8. Numoto N, Hasegawa Y, Takeda K, Miki K (2009) Inter-subunit interaction and quaternary rearrangement defined by the central stalk of prokaryotic V1-ATPase. EMBO Rep 10: 1228–1234.
- 9. Minamino T, Imada K, Namba K (2008) Mechanisms of type III protein export for bacterial flagellar assembly. Mol BioSyst 4: 1105–1115.
- 10. Minamino T, Namba K (2008) Distinct roles of the FliI ATPase and proton motive force in bacterial flagellar protein export. Nature 451: 485–488.
- 11. Minamino T, Morimoto YV, Hara N, Namba K (2011) An energy transduction mechanism used in bacterial type III protein export. Nat Commun 2: 475
- 12. Imada K, Minamino T, Tahara A, Namba K (2007) Structural similarity between the flagellar type III ATPase FliI and F1-ATPase subunits. Proc Natl Acad Sci U S A 104: 485–490.
- 13. Ibuki T, Imada K, Minamino T, Kato T, Miyata T, et al. (2011) Common architecture of the flagellar type III protein export apparatus and F- and V-type ATPases. Nat Struct Mol Biol 18: 277–282.
- 14. Lorenzini E, Singer A, Singh B, Lam R, Skarina T, et al. (2010) Structure and protein-protein interaction studies on Chlamydia trachomatis protein CT670 (YscO Homolog). J Bacteriol 192: 2746–2756.
- 15. Imamura H, Funamoto S, Yoshida M, Yokoyama K (2006) Reconstitution in vitro of V1 complex of Thermus thermophilus V-ATPase revealed that ATP binding to the A subunit is crucial for V1 formation. J Biol Chem 281: 38582–38591.
- 16. Yokoyama K, Muneyuki E, Amano T, Mizutani S, Yoshida M, et al. (1998) V-ATPase of Thermus thermophilus is inactivated during ATP hydrolysis but can synthesize ATP. J Biol Chem 273: 20504–20510.
- 17. Maher MJ, Akimoto S, Iwata M, Nagata K, Hori Y, et al. (2009) Crystal structure of A3B3 complex of V-ATPase from Thermus thermophilus. EMBO J 28: 3771–3779.
- 18. Nakano M, Imamura H, Toei M, Tamakoshi M, Yoshida M, et al. (2008) ATP hydrolysis and synthesis of a rotary motor V-ATPase from Thermus thermophilus. J Biol Chem 283: 20789–20796.
- 19. Iwata M, Imamura H, Stambouli E, Ikeda C, Tamakoshi M, et al. (2004) Crystal structure of a central stalk subunit C and reversible association/dissociation of vacuole-type ATPase. Proc Natl Acad Sci U S A 101: 59–64.
- 20. Imamura H, Nakano M, Noji H, Muneyuki E, Ohkuma S, et al. (2003) Evidence for rotation of V1-ATPase. Proc Natl Acad Sci U S A 100: 2312–2315.
- 21. Hayashi K, Ueno H, Iino R, Noji H (2010) Fluctuation theorem applied to F1-ATPase. Phys Rev Lett 104: 218103.
- 22. Ludtke SJ, Baldwin PR, Chiu W (1999) EMAN: Semiautomated software for high-resolution single-particle reconstructions. J Struct Biol 128: 82–97.
- 23. Ludtke SJ, Chiu W (2003) Focal pair merging for contrast enhancement of single particles. J Struct Biol 144: 73–78.
- 24. Katoh K, Misawa K, Kuma K, Miyata T (2002) MAFFT: a novel method for rapid multiple sequence alignment based on fast Fourier transform. Nucleic Acids Res 30: 3059–3066.
- 25. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, et al. (2011) MEGA5: Molecular Evolutionary Genetics Analysis using Maximum Likelihood, Evolutionary Distance, and Maximum Parsimony Methods. Mol Biol Evol 28: 2731–2739.
- 26. Arai S, Saijo S, Suzuki K, Mizutani K, Kakinuma Y, et al. (2013) Rotation mechanism of Enterococcus hirae V1-ATPase based on asymmetric crystal structures. Nature 493: 703–707.