Common Evolutionary Origin for the Rotor Domain of Rotary Atpases and Flagellar Protein Export Apparatus

Jun-ichi Kishikawa; Tatsuya Ibuki; Shuichi Nakamura; Astuko Nakanishi; Tohru Minamino; Tomoko Miyata; Keiichi Namba; Hiroki Konno; Hiroshi Ueno; Katsumi Imada; Ken Yokoyama

doi:10.1371/journal.pone.0064695

Abstract

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.

Introduction

Two types of rotary ATPases are found in biological membranes; V_oV₁(V type ATPase) and F_oF₁ (F type ATPase) [1]–[4]. Evolutionary counterparts of the eukaryotic V_oV₁ are found in most archea and some bacteria (often referred to as A-type ATPase). Both V_oV₁ and F_oF₁ 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 [5]. For example, V_oV₁ lacks a counterpart of the rotor shaft F₁-γ subunit [1], [4], [5].

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Figure 1. Structure of rotary ATPases and rotors.

(a) Schematic model of prokaryotic F_oF₁ (left) and V_oV₁ (right). Rotor subunits (D, F, V_o-d in V_oV₁, γ and ε in F_oF₁) are presented in brown. Rotor rings composed of hydrophobic c subunit are presented in dark brown. Peripheral stators (EG and V_o-a in V_oV₁, b₂, δ, and F_o-a in F_oF₁) 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 V_oV₁. (b) Crystal structure of γ subunit from bovine F_oF₁ (PDB: 1E79). (c) Crystal structure of DF subunit of E.hirae V_oV₁ (PDB: 3AON). (d) Crystal structure of FliJ of Salmonella typhimurium (PDB:3AJW).

https://doi.org/10.1371/journal.pone.0064695.g001

The minimal ATP-driven rotary unit of F_oF₁ 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 V_oV₁, the minimal rotary unit consists of the A₃B₃D subunits [6]. 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 [5]. However, F_oF₁ lacks a counterpart of the V₁-F subunit [1], [4], [5], which has a typical globular α/β fold [7] (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 [8].

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 [5]. 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) [9]. 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 [10], [11], 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 [12]. FliJ, which is a soluble export component protein, also shows a striking structural similarity to the coiled-coil region of F₁-γ [13]. 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 [14]. Interestingly, FliJ promotes the formation of a homo-hexamer of FliI by binding to the center of the ring, facilitating the FliI ATPase activity [9]. A very similar arrangement is observed in the F_oF₁-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 V_oV₁ and F_oF₁ and the type III export system. FliJ formed chimeric complex with A₃B₃ of V_oV₁ 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.

Methods

Proteins

The A₃B₃JL and A₃B₃DF_f expression plasmids were constructed from His-tagged V₁ (A_{(His-10/C28S/S232A/T235S/C255A/C508A)3}B_(C264S)3D_(E48C/Q55C)F) as described in Figs. S3a, S4a. The A₃B₃JL (A_{(His-10/C28S/S232A/T235S/C255A/C508A)3}B_(C264S)3JL), A₃B₃DF_f, 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) [6]. 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 [18].

FliJ was expressed in BL21(DE3)pLysS and purified as described previously [13]. 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) [13]. A₃B₃ was expressed and purified as previously described [15]. 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.

Rotation experiments

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 A₃B₃DF_f 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 [20], [21]. 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 [20], [21].

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 [22]. The focal pairs were merged using FOCALPAIR [22], [23]. The defocus amplitude, envelope and noise values were determined using CTFIT [22]. Micrographs showing significant astigmatism or drift were discarded. The particle images were selected with BOXER [22] and the boxed particle images were aligned and classified using REFINE2D.PY [22].

Phylogenetic tree analysis

All sequences used in this study were aligned using the MAFFT program (http://mafft.cbrc.jp/alignment/server/) [24]. Phylogenetic and molecular evolutionary analyses were conducted using MEGA version 5 [25]. The phylogenetic tree was constructed using the Maximum-Likelihood (ML) method under the Jones-Taylor-Thornton model by using MEGA.

Results

Reconstitution of A₃B₃J

Structural sequence alignment has previously revealed that FliJ shares some amino acid conservation with F₁-γ [13]. 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 [15] 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 [16]. 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 [16]. 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.

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Figure 2. Analysis of A₃B₃J.

(a) SDS-PAGE analysis of A₃B₃J and A₃B₃DF_f. (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.

https://doi.org/10.1371/journal.pone.0064695.g002

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 [17]. 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 [8].

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Figure 3. Structure of A₃B₃ and A₃B₃J.

Averaged cryo-EM image of A₃B₃ (a) and A₃B₃J (b) are shown. Possible ribbon models of A₃B₃ (c) and A₃B₃J (d) are also indicated below each EM image.

https://doi.org/10.1371/journal.pone.0064695.g003

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.

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Figure 4. Rotary motion of FliJ in A₃B₃.

(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.

https://doi.org/10.1371/journal.pone.0064695.g004

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 [13]. 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 V_max value of the A₃B₃JL is similar to that of A₃B₃D (Fig. S3b). Using a direct observation system similar to that described for V₁ [6], 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.

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Figure 5. Rotation of A₃B₃JL and A₃B₃DF_f.

Schematic representation of the experimental system for rotation of A₃B₃JL (a) and A₃B₃DF_f (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 A₃B₃DF_f (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.

https://doi.org/10.1371/journal.pone.0064695.g005

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 DF_f. The mutant ATPase containing the fused protein (A₃B₃DF_f) was expressed in E. coli and purified to homogeneity (Fig. 2a). The A₃B₃DF_f 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 [18]. 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 DF_f (Fig. 5c). We observed rotating beads attached to the A₃B₃DF_f at 4 mM or 10 µM ATP (Fig. 5d). Stepwise rotation of the A₃B₃DF_f 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 A₃B₃DF_f confirm that the DF_f 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.

Discussion

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 [26]. 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 V_oV₁, 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.

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Figure 6. A proposed evolution process for the rotor domains.

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 V_oV₁ have conserved the ancestral form in the rotor apparatus, C subunit (V_o-d) has been replaced with F₁-ε during evolution of F_oF₁. Function of each rotor subunit was summarized in Table S1.

https://doi.org/10.1371/journal.pone.0064695.g006

V₁-F is structurally similar to CheY, a regulatory subunit of the bacterial flagellar motor, which functions to switch the direction of rotation [7]. 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 V_o sector contains the funnel shaped C subunit (V_o-d subunit), which serves as a socket for the DF rotor in V₁ [1], [19]. In contrast, F₁-γ attaches directly onto the F_o-c ring while the F₁-ε forms contacts with both the F₁-γ and F_o-c ring [1], [3] (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 V_oV₁- and F_oF₁ (see Fig. 1a). The F₁-ε, composed of an N-terminal β sandwich and a short C terminal helix [1], [3] shows neither sequence nor structural similarity to the equivalent V_o-d (prokaryotic C subunit). Assuming that V_oV₁ have conserved the ancestral form in the rotor apparatus, V_o-d has been replaced with F₁-ε during evolution of F_oF₁.

Supporting Information

Figure S1.

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.

https://doi.org/10.1371/journal.pone.0064695.s001

(DOC)

Figure S2.

(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).

https://doi.org/10.1371/journal.pone.0064695.s002

(DOC)

Figure S3.

(a) Construction of the A₃B₃JL expression vector. (b) ATP hydrolysis activity of A₃B₃JL at the indicated [ATP].

https://doi.org/10.1371/journal.pone.0064695.s003

(DOC)

Figure S4.

(a) Construction of A₃B₃DF_f expression vector. (b) ATP hydrolysis acitivity of A₃B₃DF_f at the indicated [ATP].

https://doi.org/10.1371/journal.pone.0064695.s004

(DOC)

Figure S5.

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.

https://doi.org/10.1371/journal.pone.0064695.s005

(DOC)

Table S1.

Structure and function of each rotor subunit of Flagellar protein export apparatus, V_oV₁, and F_oF₁.

https://doi.org/10.1371/journal.pone.0064695.s006

(DOC)

Acknowledgments

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.

Author Contributions

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.

References

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Google Scholar

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Google Scholar

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Figures

Abstract

Introduction

Methods

Proteins

Rotation experiments

Electron cryo-microscopy and image analysis

Phylogenetic tree analysis

Results

Reconstitution of A3B3J

Electron cryo-microscopy analysis of A3B3J

Rotary motion of FliJ in A3B3 without uni-directionality

Anti-parallel coiled coil structure (FliJ structure) functions as a rotor

F1-γ like rotor derived from V1-D and V1-F

Discussion

Supporting Information

Figure S1.

Figure S2.

Figure S3.

Figure S4.

Figure S5.

Table S1.

Acknowledgments

Author Contributions

References

Reconstitution of A₃B₃J

Electron cryo-microscopy analysis of A₃B₃J

Rotary motion of FliJ in A₃B₃ without uni-directionality

F₁-γ like rotor derived from V₁-D and V₁-F