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F0F1-ATPase/Synthase Is Geared to the Synthesis Mode by Conformational Rearrangement of ϵ Subunit in Response to Proton Motive Force and ADP/ATP Balance*

  • Toshiharu Suzuki
    Affiliations
    ATP System Project, Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Corporation (JST), Nagatsuta 5800-2, Yokohama 226-0026, Japan

    Chemical Resources Laboratory, Tokyo Institute of Technology, Nagatsuta 4259, Yokohama 226-8503, Japan
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  • Tomoe Murakami
    Affiliations
    ATP System Project, Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Corporation (JST), Nagatsuta 5800-2, Yokohama 226-0026, Japan
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  • Ryota Iino
    Affiliations
    ATP System Project, Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Corporation (JST), Nagatsuta 5800-2, Yokohama 226-0026, Japan

    Chemical Resources Laboratory, Tokyo Institute of Technology, Nagatsuta 4259, Yokohama 226-8503, Japan
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  • Junko Suzuki
    Affiliations
    ATP System Project, Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Corporation (JST), Nagatsuta 5800-2, Yokohama 226-0026, Japan
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  • Sakurako Ono
    Affiliations
    Chemical Resources Laboratory, Tokyo Institute of Technology, Nagatsuta 4259, Yokohama 226-8503, Japan
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  • Yasuo Shirakihara
    Affiliations
    National Institute of Genetics, Mishima, Shizuoka 411-8540, Japan
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  • Masasuke Yoshida
    Correspondence
    To whom all correspondence should be addressed. Tel.: 81-45-924-5233; Fax: 81-45-924-5277;
    Affiliations
    ATP System Project, Exploratory Research for Advanced Technology (ERATO), Japan Science and Technology Corporation (JST), Nagatsuta 5800-2, Yokohama 226-0026, Japan

    Chemical Resources Laboratory, Tokyo Institute of Technology, Nagatsuta 4259, Yokohama 226-8503, Japan
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  • Author Footnotes
    * This work was supported in part by Human Frontiers Science Program Organization Grant RG15/1998-M. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Open AccessPublished:July 24, 2003DOI:https://doi.org/10.1074/jbc.M307165200
      The ϵ subunit in F0F1-ATPase/synthase undergoes drastic conformational rearrangement, which involves the transition of two C-terminal helices between a hairpin “down”-state and an extended “up”-state, and the enzyme with the up-fixed ϵ cannot catalyze ATP hydrolysis but can catalyze ATP synthesis (Tsunoda, S. P., Rodgers, A. J. W., Aggeler, R., Wilce, M. C. J., Yoshida, M., and Capaldi, R. A. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 6560–6564). Here, using cross-linking between introduced cysteine residues as a probe, we have investigated the causes of the transition. Our findings are as follows. (i) In the up-state, the two helices of ϵ are fully extended to insert the C terminus into a deeper position in the central cavity of F1 than was thought previously. (ii) Without a nucleotide, ϵ is in the up-state. ATP induces the transition to the down-state, and ADP counteracts the action of ATP. (iii) Conversely, the enzyme with the down-state ϵ can bind an ATP analogue, 2′,3′-O-(2,4,6-trinitrophenyl)-ATP, much faster than the enzyme with the up-state ϵ. (iv) Proton motive force stabilizes the up-state. Thus, responding to the increase of proton motive force and ADP, F0F1-ATPase/synthase would transform the ϵ subunit into the up-state conformation and change gear to the mode for ATP synthesis.
      F0F1-ATPase/synthase (F0F1)
      The abbreviations used are: F0F1
      the general name of F0F1-ATPase/synthase or the particular F0F1 from thermophilic Bacillus PS3, per context
      F1
      the general name of F1-ATPase or the particular F1 from thermophilic Bacillus PS3, per context
      DTNB
      5,5′-dithiobis-(2-nitro)benzoic acid
      DTT
      dithiothreitol
      FCCP
      carbonyl cyanide p-trifluoromethoxyphenylhydrazone
      TNP-AT(D)P
      2′,3′-O-(2,4,6-trinitrophenyl)-AT(D)P.
      1The abbreviations used are: F0F1
      the general name of F0F1-ATPase/synthase or the particular F0F1 from thermophilic Bacillus PS3, per context
      F1
      the general name of F1-ATPase or the particular F1 from thermophilic Bacillus PS3, per context
      DTNB
      5,5′-dithiobis-(2-nitro)benzoic acid
      DTT
      dithiothreitol
      FCCP
      carbonyl cyanide p-trifluoromethoxyphenylhydrazone
      TNP-AT(D)P
      2′,3′-O-(2,4,6-trinitrophenyl)-AT(D)P.
      catalyzes ATP synthesis/hydrolysis coupled with a transmembrane H+(proton)-translocation in bacteria, chloroplasts, and mitochondria (
      • Boyer P.D.
      ,
      • Yoshida M.
      • Muneyuki E.
      • Hisabori T.
      ,
      • Pedersen P.L.
      ,
      • Capaldi R.A.
      • Aggeler R.
      ,
      • Senior A.E.
      • Nadanaciva S.
      • Weber J.
      ). The enzyme is composed of two portions, i.e. a water-soluble F1, which has catalytic sites for ATP synthesis/hydrolysis, and a membrane-integrated F0, which mediates proton translocation. The bacterial enzyme has the simplest subunit structure, α3β3γ1δ1ϵ1 for F1 and a1b2c10–11(?) for F0. F1 is reversibly detached from F0 and is by itself a rotary motor driven by ATP hydrolysis (
      • Duncan T.M.
      • Bulygin V.V.
      • Zhou Y.
      • Hutcheon M.L.
      • Cross R.L.
      ,
      • Noji H.
      • Yasuda R.
      • Yoshida M.
      • Kinosita Jr., K.
      ,
      • Ren H.
      • Allison W.S.
      ) in which a central stalk made of γ and ϵ subunits rotates relative to the surrounding α3β3 hexamer ring where hydrolysis occurs (
      • Aggeler R.
      • Ogilvie I.
      • Capaldi R.A.
      ,
      • Schulenberg B.
      • Wellmer F.
      • Lill H.
      • Junge W.
      • Engelbrecht S.
      ). The remaining F0 portion in the membrane acts as a proton channel that mediates passive proton translocation across the membrane (
      • Fillingame R.H.
      • Dmitriev O.Y.
      ).
      The ϵ subunit is known as an endogenous inhibitor of ATPase activity of F1 and F0F1 (
      • Laget P.P.
      • Smith J.B.
      ,
      • Sternweis P.C.
      • Smith J.B.
      ). Structures of the isolated ϵ from Escherichia coli, as determined by x-ray crystallography (
      • Uhlin U.
      • Cox G.B.
      • Guss J.M.
      ) and NMR spectroscopy (
      • Wilkens S.
      • Dahlquist F.W.
      • McIntosh L.P.
      • Donaldson L.W.
      • Capaldi R.A.
      ,
      • Wilkens S.
      • Capaldi R.A.
      ), show that ϵ consists of two distinct domains. A C-terminal helical hairpin domain of ∼50 residues lies on an N-terminal 10-stranded β sandwich domain of ∼80 residues (Fig. 1A). The δ subunit (equivalent to the bacterial ϵ subunit) in the crystal structure of bovine mitochondrial F1 also has a two-domain conformation that is very similar to that of the isolated bacterial ϵ and is associated with the “bottom” globular part of the γ subunit (we refer to this conformational state of ϵ as the “down”-state hereafter) (Fig. 1B) (
      • Gibbons C.
      • Montgomery M.G.
      • Leslie A.G.W.
      • Walker J.E.
      ). However, the down-state ϵ does not exhibit an inhibitory effect on ATPase activity because, when down-state conformation is locked by cross-linking between the two domains, the inhibitory effect of ϵ is lost, and apparent activation of ATP hydrolysis is observed (
      • Schulenberg B.
      • Capaldi R.A.
      ,
      • Kato-Yamada Y.
      • Yoshida M.
      • Hisabori T.
      ). Actually, in the structure of mitochondrial F1, the δ subunit does not have any contact with the α3β3 (Fig. 1B). Another conformation of ϵ was suggested from observations that the residue (ϵSer108, E. coli numbering) in the C-terminal domain of the E. coli ϵ subunit has interactions with the residues (βGlu381) in the “DELSEED” region of the β subunit (and the homologous region of the α subunit) (
      • Aggeler R.
      • Ogilvie I.
      • Capaldi R.A.
      ,
      • Aggeler R.
      • Capaldi R.A.
      ,
      • Aggeler R.
      • Chicas-Cruz K.
      • Cai S.X.
      • Keana J.F.W.
      • Capaldi R.A.
      ,
      • Grüber G.
      • Capaldi R.A.
      ). Also, it was shown that positive residues in the C-terminal domain of the ϵ subunit of thermophilic F1 from thermophilic Bacillus PS3 would make electrostatic interaction with the DELSEED region of the β subunit (
      • Hara K.Y.
      • Kato-Yamada Y.
      • Kikuchi Y.
      • Hisabori T.
      • Yoshida M.
      ). The dynamic and flexible nature of the ϵ subunit has been also reported for chloroplast F0F1 (
      • Komatsu-Takaki M.
      ). In accordance with these biochemical results, a new conformation of ϵ was found in the crystal structure of the complex of truncated-γ (γ′) and ϵ of E. coli F1 (Fig. 1C) (
      • Rodgers A.J.
      • Wilce M.C.
      ). In this γ′ϵ complex, a helical hairpin in the previous structures of ϵ is opened, and the helices are lifted up. Such a location of the ϵ subunit could be an obstacle for the rotation of the γ subunit and, indeed, F0F1, with the ϵ locked to this lifted-up conformation by γ-ϵ cross-linking, did not show ATP hydrolysis activity. Interestingly, however, the activity of ATP synthesis of this cross-linked enzyme was fully retained (
      • Tsunoda S.P.
      • Rodgers A.J.W.
      • Aggeler R.
      • Wilce M.C.J.
      • Yoshida M.
      • Capaldi R.A.
      ). Thus, it has been established that ϵ can adopt at least two conformational states, i.e. the down-state in which C-terminal helices form a hairpin and the up-state in which the helices are extended. Only the ϵ subunit in the up-state can exert an inhibitory effect on ATPase activity.
      Figure thumbnail gr1
      Fig. 1Conformations of the ϵ subunit.A, crystal structure of the isolated E. coli ϵ subunit (
      • Wilkens S.
      • Capaldi R.A.
      ). N-terminal and C-terminal domains were shown with green and red/yellow colors, respectively. B, crystal structure of the down-state conformation of the δ subunit (equivalent to the ϵ subunit in bacterial F1) observed in bovine mitochondrial F1 (
      • Gibbons C.
      • Montgomery M.G.
      • Leslie A.G.W.
      • Walker J.E.
      ). Only subunits of βTP, γ, δ, and ϵ (no equivalent subunit in bacterial F1) are depicted in the figure. A loop that contains a DELSEED sequence was colored purple. C, crystal structure of the γ′ϵ complex of E. coli F1 (
      • Rodgers A.J.
      • Wilce M.C.
      ) superimposed with the βTP of bovine F1. Blue spheres indicate βE395 residue of the DELSEED region (second Glu residue). Cα distance between βGlu381 and ϵSer108 is 24–27 Å, too far to be cross-linked. D–F, schematic diagrams of cross-link formation in γcϵc-F1 (D), ϵcc-F1 (E), and γcϵcc-F1 (F). In the up-state, ϵCys134-γS3C is to be cross-linked. In the down-state, ϵCys134-ϵA85C is to be cross-linked. These figures were prepared by using a program package, MOLMOL (
      • Koradi R.
      • Billerter M.
      • Wuthrich K.
      ). ϵS110, ϵSer110; βE395, βGlu395; ϵC134, ϵCys134.
      Although the importance of the conformational transition of ϵ has been thus recognized, the following critical questions on this transition remain unanswered. (i) What is the actual up-state conformation of ϵ in native F0F1? The present knowledge on the up-state conformation of ϵ is largely based on the crystal structure of the γ′ϵ complex. However, it is obvious that truncated γ′ imposes an artificial constraint on the conformation of ϵ (as well as γ) in the γ′ϵ structure. Indeed, if the extreme C-terminal helix of ϵ were to have the same conformation as in the γ′ϵ, it would clash sterically with the closest β subunit. In addition, in the model reconstituted from the α3β3γ part of the mitochondrial F1 structure and the γ′ϵ structure, ϵSer108 in the γ′ϵ is apparently too far from βGlu381 to account for efficient cross-linking (Fig. 1C). In a 4.4-Å resolution electron density map of E. coli F1, the first α helix of the ϵ subunit in the extended conformation was barely seen as continuous density, but the second α helix was unable to be traced (
      • Hausrath A.C.
      • Capaldi R.A.
      • Matthews B.W.
      ). Therefore, the conformation and arrangement of the up-state ϵ in intact F0F1 is yet unclear. (ii) What is the effect of ATP and ADP on the conformational transition of ϵ in F0F1? In E. coli F1, depending on whether the added nucleotide is ATP or ADP, the same residue of the ϵE108C changes the cross-linking partner subunit; ϵ-α in Mg2+ + ATP state (in the presence of MgCl2 + 5′-adenylyl-β,γ-imidodiphosphate) and ϵ-β in the Mg2+ + ADP state (
      • Aggeler R.
      • Capaldi R.A.
      ,
      • Aggeler R.
      • Haughton M.A.
      • Capaldi R.A.
      ). However, the individual roles of ATP and ADP were not obvious for F1 from thermophilic Bacillus PS3 in our previous paper (
      • Kato-Yamada Y.
      • Yoshida M.
      • Hisabori T.
      ). The distinct role of ATP and ADP in the conformational transition of the ϵ must be clarified. (iii) Do the enzyme with the up-state ϵ and the enzyme with the down-state ϵ have different affinities to ATP and ADP? If ATP and ADP have different effects on the conformational transition of ϵ, binding affinity to ATP and ADP, conversely, might be different between the enzyme with the up-state ϵ and the enzyme with the down-state ϵ. (iv) Does the proton motive force affect the transition of ϵ? Because the enzyme with the up-state ϵ can apparently catalyze ATP synthesis but not ATP hydrolysis, the enzyme with the up-state ϵ can be regarded as the enzyme species geared to the ATP synthesis mode. If so, it is natural to expect that proton motive force would facilitate the down-to-up transition of the ϵ subunit. To address these questions, we generated a new set of F0F1 mutants from thermophilic Bacillus PS3 that enabled us to detect and fix the down- and up-states of ϵ in the working enzyme.

      EXPERIMENTAL PROCEDURES

      Preparation of the Enzymes—Plasmids for three F0F1 mutants, γcϵc-F0F1 (γS3C, ϵCys134), γcϵcc-F0F1 (γS3C, ϵA85C, ϵCys134), and ϵcc-F0F1 (ϵA85C, ϵCys134), were constructed from the plasmid pTR19-ASDS (
      • Suzuki T.
      • Ueno H.
      • Mitome N.
      • Suzuki J.
      • Yoshida M.
      ) by the Mega-primer method (
      • Landt O.
      • Grunert H.P.
      • Hahn U.
      ). Sequences of the regions amplified by PCR were verified by nucleotide sequencing. These plasmids were used for transformation of an E. coli strain DK8 (bglR, thi-1, rel-1, HfrPO1, Δ(uncB-uncC), ilv:Tn10) that lacked whole F0F1 genes. F13β3γδϵ complex) of thermophilic Bacillus strain PS3 was purified as follows. E. coli cells (DK8/pTR19-ASDS) expressing thermophilic F0F1 were disrupted in PA3-buffer (10 mm HEPES/KOH, pH 7.5, 5 mm MgCl2, and 10% glycerol) (
      • Suzuki T.
      • Ueno H.
      • Mitome N.
      • Suzuki J.
      • Yoshida M.
      ), and the cytosol fraction was obtained by removing a membrane fraction with centrifugation (150,000 × g for 20 min). The supernatant was incubated at 67 °C for 15 min, and aggregated E. coli proteins were removed by centrifugation (150,000 × g for 20 min). A yellow supernatant was supplemented with 2 volumes of 20 mm KPi buffer (pH 7.5) containing 100 mm KCl and 50 mm imidazole and applied on a nickel-nitrilotriacetic acid Superflow column (Qiagen, Hilden, Germany) equilibrated with the same buffer. After washing the column with 10 volumes of the buffer, F1 was eluted with 20 mm KPi buffer, pH 7.5, containing 100 mm KCl and 200 mm imidazole, and DTT (final concentration, 50 mm) was added to the eluted fraction. After incubation for 60 min at 25 °C, ammonium sulfate was added to the solution (final concentration, 1 m), and the solution was applied to a phenyl-Toyopearl 650 m column (Tosoh, Tokyo, Japan) equilibrated with 20 mm KPi buffer, pH 7.5, containing 0.5 mm EDTA and 1 m ammonium sulfate. The column was washed with 20 volumes of 100 mm KPi buffer, pH 7.5, containing 4 mm EDTA and 1 m ammonium sulfate to remove endogenously bound nucleotides, and a linear reverse gradient of ammonium sulfate (1–0 M) was applied. Fractions containing F1 were collected, precipitated with ammonium sulfate, and further purified with a Superdex 200HR column (Amersham Biosciences) in 20 mm HEPES/KOH buffer, pH 7.5, containing 100 mm KCl. The purified protein was frozen with liquid N2 and stored at–80 °C until use. The purified γcϵc-F1, γcϵcc-F1 and ϵcc-F1 contained 0.096 ± 0.01, 0.18 ± 0.02, and 0.18 ± 0.02 mol of ADP per mol of F1, respectively.
      Assays of Membrane Vesicles—Inverted membrane vesicles from E. coli cells expressing thermophilic F0F1 were prepared by the procedures described previously (
      • Suzuki T.
      • Ueno H.
      • Mitome N.
      • Suzuki J.
      • Yoshida M.
      ) except for a modification in which 5 mm DTT was supplemented to the cell extract just after disruption of the cells. The thermophilic F0F1 used in this work has a histidine tag of 10 residues at the N terminus of the β subunit. Prior to use, the membrane vesicles were washed twice with PA3 buffer to remove DTT. ATPase activities of the membrane vesicles containing the F0F1 mutants were inactivated by N,N′-dicyclohexylcarbodiimide down to < 20% of the initial activities, which were almost the same as in the case of the wild-type (15–20%). ATP-driven proton pump activity of membrane vesicles was assayed with fluorescence quenching of 9-amino-6-chloro-2-methoxyacridine at 40 °C in PA4 buffer (10 mm HEPES/KOH, pH 7.5, 100 mm KCl, and 5 mm MgCl2) as described previously (
      • Suzuki T.
      • Ueno H.
      • Mitome N.
      • Suzuki J.
      • Yoshida M.
      ). The reaction was started by the addition of 1 mm ATP and terminated by the addition of 1 μm carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP). ATP synthesis activity was measured at 50 °C in PA4-buffer containing 1 mm ADP, 25 mm KPi, pH 7.5, and membrane vesicles (4.5 μg of protein/ml). Oxidized and reduced membrane vesicles were prepared by treating with 20 μm CuCl2 for 30 min and 10 mm DTT for 30 min, respectively. EDTA (final concentration, 1 mm) was added to the oxidized vesicle solution prior to the assay of ATP synthesis to chelate free Cu2+. EDTA was also added to the reduced vesicle solution to adjust the conditions. After a 5-min preincubation, the reaction was initiated by adding 5 mm NADH and terminated at 2, 4, 6, 8, 10, and 12 min by adding 2.5% trichloroacetic acid. The solution was neutralized to pH 7.7 with 0.25 m Tris acetate (pH 9.5), and the amount of synthesized ATP was determined with ATP bioluminescence assay kit CLSII (Roche Applied Science).
      Other Assays—ATPase activity was monitored in triplicate in 50 mm HEPES/KOH, pH 7.5, containing 100 mm KCl, 5 mm MgCl2, and 3 mm ATP with an ATP-regenerating system (
      • Suzuki T.
      • Suzuki J.
      • Mitome N.
      • Ueno H.
      • Yoshida M.
      ), and average hydrolysis rates in a time period from 3 to 6 min after initiation of the reactions at 40 °C were measured. The activity that hydrolyzed 1 μmol of ATP per minute was defined as one unit. 2′,3′-O-(2,4,6-trinitrophenyl)-ATP (TNP-ATP) and 2′,3′-O-(2,4,6-trinitrophenyl)-ADP (TNP-ADP) were purchased from Molecular Probes (Eugene, OR). Fluorescence change induced by the binding of TNP-nucleotide to the enzyme was monitored in a Spectrofluorometer model FP-6500 (Jasco, Tokyo, Japan) as performed previously (
      • Kato Y.
      • Matsui T.
      • Tanaka N.
      • Muneyuki E.
      • Hisabori T.
      • Yoshida M.
      ). Protein concentrations were determined by using the BCA protein assay kit from Pierce, with bovine serum albumin as a standard.

      RESULTS

      Mutants—We generated three mutants. To obtain the enzyme with the up-fixed ϵ by cross-linking between γ and ϵ, one cysteine residue was introduced into the N-terminal region of the γ subunit (γS3C) and another was added to the C-terminal end of ϵ subunit (ϵCys134) (Fig. 1D). To obtain the enzyme with the down-fixed ϵ by cross-linking two domains within ϵ, a mutant that had ϵA85C and ϵCys134 was used (Fig. 1E). To assess the relative population of enzymes with up- and down-state ϵ under various conditions, three mutations, γS3C, ϵA85C and ϵCys134 were introduced. The ϵAla85 is located in a region between two domains of ϵ and abuts on ϵCys134 (Cα distance is 9.0 Å) when the C-terminal domain adopts a hairpin structure (
      • Uhlin U.
      • Cox G.B.
      • Guss J.M.
      ). Therefore, ϵCys134 is expected to make the cross-link with γS3C when the ϵ is in the up-state or with ϵA85C when the ϵ is in the down-state (Fig. 1F). These mutations were termed γcϵc, ϵcc, and γcϵcc, respectively. The enzymes have one endogenous cysteine residue in the F0a subunit. This cysteine is buried inside of the transmembrane region and does not respond to the CuCl2 and 5,5′-dithiobis-(2-nitro)benzoic acid (DTNB) treatment employed in this report. The enzymes containing γcϵc, ϵcc, and γcϵcc were as active as the wild-type enzyme in their reduced forms (Table I). Also ATPase activities of the inverted membrane vesicles prepared from the cells expressing the wild-type and the F0F1 mutants were similar to each other under the reducing conditions. SDS-PAGE analysis of the membrane vesicles showed almost identical band patterns for the three mutants and the wild-type (not shown). Therefore, the amounts of expressed mutant F0F1 in the inverted membranes are similar to that of the wild-type F0F1.
      Table IATPase activity of the wild-type and mutant F1 and F0F1
      Enzymes
      Reduced form of the enzymes was used for the analysis.
      ATPase activity
      ATPase activity was determined at 40 °C in the presence of 3 mm ATP. Values for F0F1 are the activity per 1 mg of membrane proteins.
      units/mg/min
      Purified F1
       Wild-type10.6 ± 0.1
       γcϵc9.4 ± 0.1
       γcϵcc11.3 ± 0.5
       ϵcc9.9 ± 0.2
      F0F1 (membrane vesicles)
       Wild-type0.99 ± 0.02
       γcϵc0.89 ± 0.03
       γcϵcc1 ± 0.02
       ϵcc1 ± 0.1
      a Reduced form of the enzymes was used for the analysis.
      b ATPase activity was determined at 40 °C in the presence of 3 mm ATP. Values for F0F1 are the activity per 1 mg of membrane proteins.
      Cross-linking of ϵ in the Up-state—In the previous experiments using E. coli F0F1 (
      • Tsunoda S.P.
      • Rodgers A.J.W.
      • Aggeler R.
      • Wilce M.C.J.
      • Yoshida M.
      • Capaldi R.A.
      ), cysteine residues were introduced at positions γ99 and ϵ118 (E. coli numbering) to fix the conformation of ϵ in the up-state by a γ-ϵ cross-link. These positions were chosen based on the crystal structure of the γ′ϵ complex in which γ99 is located in the globular domain of γ′ and ϵ118 is in the extreme C-terminal helix of ϵ. In this structure, two helices do not fully extend but rather entwine the globular domain of γ′ (Fig. 1C). Expecting that helices of the up-state ϵ in the native enzyme could extend straighter, we introduced cysteine residues to a near N-terminal position of γ (γSer3) atop the central helical coiled-coil of the γ subunit and to the C terminus of ϵ (ϵ134) (Fig. 1D). The membrane vesicles of E. coli expressing γcϵc-F0F1 were oxidized in 20 μm CuCl2 for 20 min at 25 °C and analyzed with non-reducing SDS-PAGE (SDS-PAGE without prior reducing treatment) (Fig. 2A). Compared with a control γcϵc-F0F1, which was treated with 50 mm DTT prior to electrophoresis (Fig. 2A, lane 1), a new band appeared just below the band of the β subunit (Fig. 2A, lane 4, indicated by an arrow). Peptide sequencing of this band gave two kinds of amino acid sequences corresponding to the N terminus sequences of γ and ϵ, indicating that this band is a cross-link product of these two subunits. Consistently, band intensities of γ and ϵ decreased. The same γ-ϵ cross-link product was also readily generated in the purified γcϵc-F1 under the same oxidizing conditions (Fig. 2A, lane 5). Cross-linking yields in the F1 and F0F1 were estimated from the band intensities to be 80–85%. It is worth noting that γ-ϵ cross-link was generated spontaneously in ∼40% of F1 during the purification (2 days) that was carried out without DTT and EDTA. Also, ∼60% of F0F1 in membrane vesicles were spontaneously oxidized during preparation. The efficient cross-linking between γS3C and ϵCys134 suggests that their proximal location is in the up-state conformation of ϵ in F1 and F0F1 and that the C-terminal helix of ϵ inserts itself deep into the central cavity of the α3β3. Because the isolated γcϵc-F1 used in the above experiments was mostly free from endogenous nucleotide, the ϵ mostly adopts the up-state in the absence of bound nucleotides.
      Figure thumbnail gr2
      Fig. 2Cross-link between N terminus of the γ subunit and the C terminus of the ϵ subunit.A, non-reducing SDS-PAGE analysis. Lanes 1 and 4, membrane vesicles containing γcϵc-F0F1; lanes 2 and 5, isolated γcϵc-F1; lane 3, isolated wild-type F0F1 (MK). All samples except for lane 3 were treated with 20 μm CuCl2 for 30 min at 25 °C. Then, samples of lanes 1 and 2 were reduced with 50 mm DTT for 1 h. A band of γ-ϵ cross-linked product appeared just below the band of the β subunit. B, effect of the γ-ϵ cross-linking on ATPase activities of the isolated γcϵc-F1 (left panel) and membrane vesicles containing γcϵc-F0F1 (right panel). ATP hydrolysis by the reduced (white bars) and oxidized (black bars) γcϵc-F1 and membrane vesicles containing γcϵc-F0F1 were assayed at 40 °C. The same procedures were applied to the wild-type F1 and F0F1. C, effect of the γ-ϵ cross-linking on proton pump activity. Proton pump activities of the reduced or oxidized membrane vesicles were analyzed by monitoring the fluorescence of 9-amino-6-chloro-2-methoxyacridine at 40 °C. Prior to the analysis, 1 mm EDTA was added to the solutions to remove free Cu2+. At the indicated times, pumping was initiated by adding 1 mm ATP and terminated by 1 μg/ml FCCP. D, effect of the γ-ϵ cross-linking on ATP synthesis activity. The reactions were started by addition of 5 mm NADH to the membrane vesicle solutions containing reduced or oxidized wild-type F0F1 and γcϵc-F0F1. The reactions were carried out at 50 °C, and the amount of generated ATP was measured with luciferase.
      Activities of F1 and F0F1 with the Up-fixed ϵ The ATPase activity of γcϵc-F1 was severely inhibited by oxidation with its residual activity being only 21% of that of the reduced γcϵc-F1, whereas the ATPase activity of the wild-type F1 was hardly affected whether it was oxidized or reduced (Fig. 2B, left panel). The degree of inhibition by oxidation (79%) agreed well with the yield of cross-link by oxidation (81%). Similarly, the ATPase activity of the γcϵc-F0F1 contained in the vesicles was inhibited (77%) in proportion to the yield of cross-linking (80%) (Fig. 2B, right panel). Because ATP hydrolysis was blocked, oxidized γcϵc-F0F1 was unable to mediate ATP-driven proton translocation, whereas reduced γcϵc-F0F1 was fully capable of it (Fig. 2C). ATP synthesis activities of membrane vesicles containing reduced or oxidized γcϵc-F0F1 were also measured. Membrane vesicles containing the wild-type and γcϵc-F0F1 treated with DTT catalyzed ATP synthesis at 44.5 ± 2.8 and 34.7 ± 2.4 nmol of ATP/min/mg of membrane protein, respectively. Oxidized vesicles showed 71% (wild-type) and 75% (γcϵc-F0F1) of the ATP synthesis activity of the vesicles treated with DTT (Fig. 2D). Thus, ATP synthesis activity was retained after the formation of the γ-ϵ cross-link to lock the ϵ in the up-state. These results are consistent with the previous reports, demonstrating remarkable asymmetric inhibition by the up-state ϵ toward ATP hydrolysis (
      • Schulenberg B.
      • Capaldi R.A.
      ,
      • Tsunoda S.P.
      • Rodgers A.J.W.
      • Aggeler R.
      • Wilce M.C.J.
      • Yoshida M.
      • Capaldi R.A.
      ,
      • Nowak K.F.
      • Tabidze V.
      • McCarty R.E.
      ).
      Effect of ATP and ADP on the Conformational State of ϵ To assess the distribution of the ϵ either in the up-state or down-state, γcϵcc-F1 and γcϵcc-F0F1 were used. With oxidation procedures, the down-state ϵ can be detected as a band corresponding to an internally cross-linked ϵ (ϵA85C and ϵC134) and the up-state ϵ as a γ-ϵ band (γS3C and ϵCys134). The ϵ subunit behaved very similarly in γcϵcc-F1 and γcϵcc-F0F1 (Fig. 3, A and B). In the absence of nucleotides, the ϵ in F1 and in F0F1 was mostly in the up-state (Fig. 3, A and B, lanes 2), and the up-state conformation was stabilized when 3 mm ADP was present (Fig. 3, A and B, lanes 3).
      We reported previously that ADP, though less effective than ATP, induced the down-state conformation of the ϵ subunit in the α3β3γϵ complex (
      • Kato-Yamada Y.
      • Yoshida M.
      • Hisabori T.
      ). When we used ADP pretreated with hexokinase and glucose in the experiment, the down-state conformation was not detected. Therefore, contaminated ATP in the commercial ADP might be the reason for the previous result.
      The further addition of 5 mm Pi caused no significant change (not shown). However, the γ-ϵ band disappeared, and the internally cross-linked ϵ band (Fig. 3, A and B, arrowheads) appeared when ADP was converted into ATP by pyruvate kinase (Fig. 3, A and B, lanes 4). Also, the internally cross-linked ϵ band appeared when ATP was added from the beginning (Fig. 3, A and B, lanes 5). The addition of hexokinase and glucose to the sample of the lanes 5 (Fig. 3, A and B) resulted in the appearance of the γ-ϵ band (Fig. 3, A and B, lanes 6). Thus, it is clear that the ϵ subunit in F1 and F0F1 adopts reversibly the up-state conformation in the presence of ADP and the down-state conformation in the presence of ATP. As shown previously (
      • Kato-Yamada Y.
      • Yoshida M.
      • Hisabori T.
      ), hydrolysis of ATP is not necessary to stabilize the down-state ϵ, because 3 mm AMP-PNP also stabilized the down-state conformation of ϵ (not shown).
      Figure thumbnail gr3
      Fig. 3Effect of ATP and ADP on the conformational state of the ϵ subunit.A, analysis of γcϵcc-F1. γcϵcc-F1 was incubated with indicated components for 2 min. Concentrations of ADP (lanes 3 and 4) and ATP (lanes 5 and 6)were3mm. For the sample of lane 4, 125 μg/ml pyruvate kinase and 9.4 mm phosphoenolpyruvate (final concentrations) were added and incubated for 1 min. For the sample of lane 6, 9.4 units/ml hexokinase and 38 mm glucose (final concentrations) were added and incubated for 1 min. The samples were reduced with 50 mm DTT (lane 1) or oxidized with 20 μm CuCl2 (lanes 2–6) for 10 min and applied to non-reducing SDS-PAGE. Lanes 1 and 2, no nucleotide; lane 3, ADP; lane 4, ADP followed by pyruvate kinase treatment; lane 5, ATP; lane 6, ATP followed by hexokinase (Hex) treatment. Arrowheads in lanes 4 and 5 indicate the position of the ϵ subunit with internal cross-link. All reactions were carried out in 50 mm HEPES/NaOH, pH 7.5, containing 100 mm NaCl and 5 mm MgCl2 at 50 °C. A distorted band above of the α subunit band in lane 4 was pyruvate kinase. B, analysis for γcϵcc-F0F1. Membrane vesicles containing γcϵcc-F0F1 were used for the analysis. Lane 1, reduced γcϵcc-F0F1, no nucleotide; lanes 2–6, membrane vesicles containing γcϵcc-F0F1 treated with the same procedures as for panel A; lane 7, γcϵcc-F1 incubated with ATP (the same sample as in panel A, lane 5) to show the band position of the ϵ subunit with internal cross-link. The samples were reduced with 50 mm DTT (lane 1) or oxidized with 100 μm CuCl2 (lanes 2–6) and applied to non-reducing SDS-PAGE.
      TNP-AT(D)P Binding to F1 with Up- or Down-state ϵ It has been known that a nucleotide analogue, TNP-AT(D)P, increases its fluorescence upon binding to F1 (
      • Kato Y.
      • Matsui T.
      • Tanaka N.
      • Muneyuki E.
      • Hisabori T.
      • Yoshida M.
      ). Taking advantage of this, we compared initial kinetics of nucleotide binding to the enzymes that contained the up- or down-state ϵ. To measure the binding to the nucleotide binding site with the highest affinity, a sub-stoichiometric amount of TNP-AT(D)P was mixed with γcϵc-F1 or ϵcc-F1, and fluorescence changes were monitored. Time courses of TNP-ADP binding were almost the same for γcϵc-F1 and ϵcc-F1, irrespective of whether they were reduced or oxidized (Fig. 4, A and B). The wild-type F1, with or without oxidizing treatment, also showed the same kinetics of TNP-ADP binding (not shown). These results indicated that TNP-ADP binding to F1 was not affected by the conformational states of the ϵ subunit. The time course of TNP-ATP binding to reduced γcϵc-F1 was also the same as that of ϵcc-F1 (Fig. 4C) and wild-type F1 (not shown), ensuring no significant effect of the introduced cysteines on the TNP-ATP binding kinetics of F1. The time course of TNP-ATP binding to the oxidized γcϵc-F1 (Fig. 4D, bottom curve) was similar to that of TNP-ADP binding to the oxidized γcϵc-F1, indicating that TNP-ATP and TNP-ADP bind to the same site of F1 with the up-fixed ϵ. The oxidized ϵcc-F1, on the contrary, bound TNP-ATP much faster, and the fluorescence reaches a higher magnitude than with the oxidized γcϵc-F1. (Fig. 4D, upper curve). Thus, F1 with the down-state ϵ binds TNP-ATP quickly, whereas F1 with the up-state ϵ binds it slowly. Accordingly, results of TNP-ATP binding to the reduced γcϵc-F1 and ϵcc-F1 in Fig. 4C are well interpreted as a mixture of F1s with the up- and down-state ϵ.
      Figure thumbnail gr4
      Fig. 4TNP-ATP binding to F1 with the up- or down-fixed ϵ subunit. To obtain F1 with the up- or down-fixed ϵ subunit by γ-ϵ cross-linking (γcϵc-F1) or internal cross-linking within ϵ (ϵcc-F1), purified γcϵc-F1 and ϵcc-F1 in 50 mm HEPES/KOH, pH7.5, were oxidized with 20 μmcϵc-F1) or 100 μmcc-F1) CuCl2 for 1 h at 25 °C. Reduced samples were not subjected to this oxidation procedures. TNP-AT(D)P (50 nm) was incubated at 50 °C for 5 min in 50 mm HEPES/KOH, pH 7.5, containing 100 mm KCl, 5 mm MgCl2, and 20 μl of F1 (6 μm) were added into the cuvette (final F1 concentration, 100 nm) at the time indicated with arrows (final volume, 1.2 ml). Fluorescence change was monitored at 548 nm with an excitation light at 410 nm. Red traces, γcϵc-F1; blue traces, ϵcc-F1. Apparent rate constants for the nucleotide binding were estimated by fitting single exponential functions to the time courses, as shown.
      Effect of Proton Motive Force on the State of ϵ The inverted membrane vesicles containing γcϵcc-F0F1 were incubated for 3 min in the varying amounts of ATP and ADP, and conformational states of the ϵ subunit were analyzed with non-reducing SDS-PAGE after fixing the conformation by cross-linking (Fig. 5A, lanes 1–6). As ATP increased and ADP decreased, intensity of the γ-ϵ band decreased, as is expected from the results mentioned above. However, when the incubation was continued for another 5 min after the addition of NADH to impose proton motive force, the intensity of the γ-ϵ band did not significantly decrease even at high ATP concentrations (Fig. 5A, lanes 7–12). When FCCP, an uncoupler that dissipates proton motive force, was added in addition to NADH, the intensity of the γ-ϵ band was decreased as ATP increased, similar to Fig. 5A, lanes 1–6 (Fig. 5A, lanes 13–18). These results suggest that when proton motive force is provided, the ϵ subunit in F0F1 strongly favors the up-state conformation irrespective of ADP/ATP balance. In other words, proton motive force counteracts the effect of ATP in the conformational transition of the ϵ subunit.
      Figure thumbnail gr5
      Fig. 5Effect of proton motive force on the conformational state of ϵ subunit in F0F1.A, membrane vesicles containing γcϵcc-F0F1 were incubated for 3 min with mixtures of varying amounts of ATP and ADP (ATP + ADP = 3 mm). An aliquot of each reaction mixture was transferred to another tube, treated with 200 μm DTNB for 5 min to fix the conformational state of the ϵ subunit by cross-linking, and subjected to non-reducing SDS-PAGE analysis (lanes 1–6). For the remaining reaction mixtures, 5 mm NADH (lanes 13–18)or5mm NADH + 3 μg/ml FCCP (lanes 6–12) was added, and incubation was continued for 5 min. Then, the reaction mixtures were treated with 200 μm DTNB for 5 min and analyzed with non-reducing SDS-PAGE. DTNB was used instead of CuCl2 as an oxidant, because DTNB is less harmful to the membrane vesicles than CuCl2. Concentrations of ATP and ADP were as follows: 0 and3mm (lanes 1, 7, and 13); 0.3 and 2.7 mm (lanes 2, 8, and 14); 0.5 and 2.5 mm (lanes 3, 9, and 15); 1 and 2 mm (lanes 4, 10, and 16); 2 and 1 mm (lanes 5, 11, and 17); and 2.7 and 0.3 mm (lanes 6, 12, and 18), respectively. The experiments were carried out at 50 °C in 50 mm HEPES/NaOH, pH 7.5, containing 100 mm NaCl and 5 mm MgCl2. It was confirmed that NADH oxidation by respiratory chains of vesicles under these conditions was as active as that at 37 °C and could generate proton motive force. In this figure, γ-ϵ bands formed by cross-linking are shown. B, the relative staining intensities of the γ-ϵ bands in panel A were plotted against the ATP concentrations. Closed triangles, the control samples (lanes 1–6); closed circles, NADH (lanes 7–12); open circles, NADH + FCCP (lanes 13–18).

      DISCUSSION

      C Terminus of the ϵ Subunit Reaches the Center of F1—The questions listed in the Introduction were mostly answered by the present study. Concerning the first question (i), it becomes evident that C terminus of ϵ in the up-state is located near the N terminus of the γ subunit.
      Very recently, we have succeeded in determining crystal structure of an α3β3γϵ complex of thermophilic F1 (Y. Shirakihara, M. Yoshida, and T. Suzuki, unpublished results). In the structure, C-terminal helices of the ϵ subunit indeed extend straight, and the C terminus of the ϵ subunit is close to the N terminus of the γ subunit.
      To reach this position, referring to the structure of mitochondrial F1, the C-terminal helices of ϵ have to extend ∼70 Å from the exit (ϵA85) of the N-terminal β sandwich domain. Considering the length of α-helix per residue (1.5 Å/residue) (
      • Dunn S.D.
      • McLachlin D.T.
      • Revington M.
      ), a peptide stretch of 48 residues from ϵAla85 to ϵLys133 can extend by 72 Å as an α-helix or longer as two helices with a connecting segment. Previous cross-linking results of E. coli F1 between βGlu381 and ϵSer108 (
      • Aggeler R.
      • Ogilvie I.
      • Capaldi R.A.
      ,
      • Aggeler R.
      • Capaldi R.A.
      ,
      • Aggeler R.
      • Chicas-Cruz K.
      • Cai S.X.
      • Keana J.F.W.
      • Capaldi R.A.
      ,
      • Grüber G.
      • Capaldi R.A.
      ) are explained by this new arrangement rather than by the γ′ϵ structure (Fig. 1, compare C and D) (
      • Rodgers A.J.
      • Wilce M.C.
      ). Probably, the γ′ϵ structure represents an intermediate conformation that appears during the transition of ϵ from the down-state to the up-state. Our study suggests that three helices, the coiled-coil of the γ subunit and C-terminal helix of the ϵ subunit, rather than two as previously thought, rotate as a body within the α3β3 ring when F0F1 with the up-state ϵ is synthesizing ATP.
      ATP and ADP Have Opposite Effects on the Conformational States of the ϵ Subunit—As to the second question (ii), it is now clear that the ϵ subunit, either in F1 or F0F1, is in the up-state conformation in the absence of nucleotide or the presence of ADP and is in the down-state conformation in the presence of ATP. Thus, ATP and ADP counteract each other (Fig. 6). Reciprocally, an ATP analogue, TNP-ATP, binds quickly to F1 with down-state ϵ but slowly to F1 with up-state ϵ. An ADP analogue, TNP-ADP, does not show binding preference between F1s with up- and down-state ϵ. Therefore, if TNP-AT(D)P mimics the AT(D)P correctly in binding to F1, the answer to the third question (iii) will be that F1s with down-state ϵ indeed prefer ATP to ADP, whereas F1s with up-state ϵ bind both ATP and ADP in the same slow kinetics. The results are consistent with the previous observation that the α3β3γ complex binds TNP-ATP quickly, but the reconstituted α3β3γϵ complex does this slowly (
      • Kato Y.
      • Matsui T.
      • Tanaka N.
      • Muneyuki E.
      • Hisabori T.
      • Yoshida M.
      ), because without previous exposure to nucleotide, the ϵ subunit in the reconstituted α3β3γϵ must be in the up-state.
      Figure thumbnail gr6
      Fig. 6Schematic diagram of two forms of the F0F1 with up-state ϵ (left) and down-state ϵ (right). The F0F1 with up-state ϵ can catalyze ATP synthesis but not ATP hydrolysis (ATP synthesis mode). This form is stabilized by ADP and proton motive force. The F0F1 with down-state ϵ can catalyze ATP synthesis as well as ATP hydrolysis (proton pump/ATP synthesis mode). This form is favored when ATP is present. Transition between two forms is determined by proton motive force and ADP/ATP balance.
      The ϵ Subunit Transits between Two States Depending on Proton Motive Force and ADP/ATP—This study has revealed that proton motive force counteracts the effect of ATP by stabilizing the up-state ϵ (answer to the fourth question (iv)). Therefore, the two conformational states of ϵ in F0F1 are alternated by two factors, i.e. proton motive force and ADP/ATP balance (Fig. 6). At high proton motive force and low ATP, ϵ is predominantly in the up-state, and F0F1 is geared to the ATP synthesis mode. At low proton motive force and high ATP, ϵ adopts the down-state and F0F1 hydrolyzes ATP to pump out protons, generating proton motive force with enough magnitude to drive uptake of nutrients and flagella motion.
      Role of C-terminal Helices of the ϵ Subunit—In some bacteria, such as Chlorobium limicola (
      • Xie D.
      • Lill H.
      • Hauska G.
      • Maeda M.
      • Futai M.
      • Nelson N.
      ) and Thermotoga neapolitana (
      • Iida T.
      • Inatomi K.
      • Kamagata Y.
      • Maruyama T.
      ), the native ϵ subunit lacks the C-terminal helical domain. Without the C-terminal helical domain, the ϵ subunit cannot adopt the up-state arrangement and should be always in the state that is functionally similar to the down-state. These bacteria grow in anaerobic environments, and F0F1 should work as an ATP hydrolysis-driven proton pump. Because the F0F1 with up-state ϵ is unable to mediate ATP hydrolysis-driven proton pumping, these bacteria do not need, or even had better delete, the C-terminal domain of the ϵ subunit. F0F1 with down-state ϵ can catalyze both ATP synthesis and ATP hydrolysis (
      • Tsunoda S.P.
      • Rodgers A.J.W.
      • Aggeler R.
      • Wilce M.C.J.
      • Yoshida M.
      • Capaldi R.A.
      ). Therefore, it is not surprising that a mutant E. coli F0F1 containing the ϵ subunit with deleted C-terminal helical domain or an artificially fused protein at the C terminus can support aerobic growth by oxidative phosphorylation (
      • Kuki M.
      • Noumi T.
      • Maeda M.
      • Amemura A.
      • Futai M.
      ,
      • Cipriano D.J.
      • Bi Y.
      • Dunn S.D.
      ). A similar observation was reported recently for chloroplast F0F1 (
      • Nowak K.F.
      • Tabidze V.
      • McCarty R.E.
      ). Then, a critical question should be asked, i.e. what is the essential function of the F0F1 whose ϵ subunit is in the up-state? Probably, the F0F1 with the up-state ϵ plays an important role under starving conditions rather than in rich nutritional environments. In E. coli cells, total concentration of cellular adenine nucleotides is maintained to be ∼3 mm (
      • Neuhard J.
      • Nygaard P.
      ), but the fraction of ATP in total adenine nucleotide pool varies from 3 to 0.3 mm in parallel with growth rate (
      • Franzen J.S.
      • Binkley S.B.
      ,
      • Smith R.C.
      • Maaløe O.
      ,
      • Bagnara A.S.
      • Finch L.R.
      ) through ribosome synthesis (
      • Gaal T.
      • Bartlett M.S.
      • Ross W.
      • Turnbough Jr., C.L.
      • Gourse R.L.
      ) and transcription (
      • Alper S.
      • Dufour A.
      • Garsin D.A.
      • Duncan L.
      • Losick R.
      ). As ATP concentration decreases from 3 to 0.3 mm in the absence of proton motive force, the population of the F0F1 with up-state ϵ increases about 3-fold (Fig. 5B) so that hydrolysis of the precious ATP by F0F1 is suppressed. For any organisms, regulation of ATP synthesis/hydrolysis to meet physiological demand in quickly changing nutritional conditions is a critical matter, and conformational transition of the ϵ subunit in F0F1 might constitute a part of an elaborately integrated regulatory system that awaits further study.

      Acknowledgments

      We thank our colleagues, Drs. H. Ueno, N. Mitome, T. Hisabori, and T. Mogi, for helpful discussions. Analytical condition for ATP synthesis activity was optimized by the assistance of N. Mitome.

      References

        • Boyer P.D.
        Annu. Rev. Biochem. 1997; 66: 717-749
        • Yoshida M.
        • Muneyuki E.
        • Hisabori T.
        Nat. Rev. Mol. Cell Biol. 2001; 2: 669-677
        • Pedersen P.L.
        J. Bioenerg. Biomembr. 2002; 34: 327-332
        • Capaldi R.A.
        • Aggeler R.
        Trends Biochem. Sci. 2002; 27: 154-160
        • Senior A.E.
        • Nadanaciva S.
        • Weber J.
        Biochim. Biophys. Acta. 2002; 1553: 188-211
        • Duncan T.M.
        • Bulygin V.V.
        • Zhou Y.
        • Hutcheon M.L.
        • Cross R.L.
        Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 10964-10968
        • Noji H.
        • Yasuda R.
        • Yoshida M.
        • Kinosita Jr., K.
        Nature. 1997; 386: 299-302
        • Ren H.
        • Allison W.S.
        Biochim. Biophys. Acta. 2000; 1458: 221-233
        • Aggeler R.
        • Ogilvie I.
        • Capaldi R.A.
        J. Biol. Chem. 1997; 272: 19621-19624
        • Schulenberg B.
        • Wellmer F.
        • Lill H.
        • Junge W.
        • Engelbrecht S.
        Eur. J. Biochem. 1997; 249: 134-141
        • Fillingame R.H.
        • Dmitriev O.Y.
        Biochim. Biophys. Acta. 2002; 1565: 232-245
        • Laget P.P.
        • Smith J.B.
        Arch. Biochem. Biophys. 1979; 197: 83-89
        • Sternweis P.C.
        • Smith J.B.
        Biochemistry. 1980; 19: 526-531
        • Uhlin U.
        • Cox G.B.
        • Guss J.M.
        Structure. 1997; 5: 1219-1230
        • Wilkens S.
        • Dahlquist F.W.
        • McIntosh L.P.
        • Donaldson L.W.
        • Capaldi R.A.
        Nat. Struct. Biol. 1995; 2: 961-967
        • Wilkens S.
        • Capaldi R.A.
        J. Biol. Chem. 1998; 273: 26645-26651
        • Gibbons C.
        • Montgomery M.G.
        • Leslie A.G.W.
        • Walker J.E.
        Nat. Struct. Biol. 2000; 7: 1055-1061
        • Schulenberg B.
        • Capaldi R.A.
        J. Biol. Chem. 1999; 274: 28351-28355
        • Kato-Yamada Y.
        • Yoshida M.
        • Hisabori T.
        J. Biol. Chem. 2000; 275: 35746-35750
        • Aggeler R.
        • Capaldi R.A.
        J. Biol. Chem. 1996; 271: 13888-13891
        • Aggeler R.
        • Chicas-Cruz K.
        • Cai S.X.
        • Keana J.F.W.
        • Capaldi R.A.
        Biochemistry. 1992; 31: 312956-312961
        • Grüber G.
        • Capaldi R.A.
        J. Biol. Chem. 1996; 271: 32623-32628
        • Hara K.Y.
        • Kato-Yamada Y.
        • Kikuchi Y.
        • Hisabori T.
        • Yoshida M.
        J. Biol. Chem. 2001; 276: 23969-23973
        • Komatsu-Takaki M.
        Eur. J. Biochem. 1993; 214: 587-591
        • Rodgers A.J.
        • Wilce M.C.
        Nat. Struct. Biol. 2000; 7: 1051-1054
        • Tsunoda S.P.
        • Rodgers A.J.W.
        • Aggeler R.
        • Wilce M.C.J.
        • Yoshida M.
        • Capaldi R.A.
        Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 6560-6564
        • Hausrath A.C.
        • Capaldi R.A.
        • Matthews B.W.
        J. Biol. Chem. 2001; 276: 47227-47232
        • Aggeler R.
        • Haughton M.A.
        • Capaldi R.A.
        J. Biol. Chem. 1995; 270: 9185-9191
        • Suzuki T.
        • Ueno H.
        • Mitome N.
        • Suzuki J.
        • Yoshida M.
        J. Biol. Chem. 2002; 277: 13281-13285
        • Landt O.
        • Grunert H.P.
        • Hahn U.
        Gene. 1990; 96: 125-128
        • Suzuki T.
        • Suzuki J.
        • Mitome N.
        • Ueno H.
        • Yoshida M.
        J. Biol. Chem. 2000; 275: 37902-37906
        • Kato Y.
        • Matsui T.
        • Tanaka N.
        • Muneyuki E.
        • Hisabori T.
        • Yoshida M.
        J. Biol. Chem. 1997; 272: 24906-24912
        • Nowak K.F.
        • Tabidze V.
        • McCarty R.E.
        Biochemistry. 2002; 41: 15130-15134
        • Dunn S.D.
        • McLachlin D.T.
        • Revington M.
        Biochim. Biophys. Acta. 2000; 1458: 356-363
        • Xie D.
        • Lill H.
        • Hauska G.
        • Maeda M.
        • Futai M.
        • Nelson N.
        Biochim. Biophys. Acta. 1993; 1172: 267-273
        • Iida T.
        • Inatomi K.
        • Kamagata Y.
        • Maruyama T.
        Extremophiles. 2002; 6: 369-375
        • Kuki M.
        • Noumi T.
        • Maeda M.
        • Amemura A.
        • Futai M.
        J. Biol. Chem. 1988; 263: 17437-17442
        • Cipriano D.J.
        • Bi Y.
        • Dunn S.D.
        J. Biol. Chem. 2002; 277: 16782-16790
        • Neuhard J.
        • Nygaard P.
        Neidhardt F.C. Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology. 1st Ed. ASM Press, Washington, D. C.1987: 445-473
        • Franzen J.S.
        • Binkley S.B.
        J. Biol. Chem. 1961; 236: 515-519
        • Smith R.C.
        • Maaløe O.
        Biochim. Biophys. Acta. 1964; 86: 229-234
        • Bagnara A.S.
        • Finch L.R.
        Eur. J. Biochem. 1973; 36: 422-427
        • Gaal T.
        • Bartlett M.S.
        • Ross W.
        • Turnbough Jr., C.L.
        • Gourse R.L.
        Science. 1997; 278: 2092-2097
        • Alper S.
        • Dufour A.
        • Garsin D.A.
        • Duncan L.
        • Losick R.
        J. Mol. Biol. 1996; 260: 165-177
        • Koradi R.
        • Billerter M.
        • Wuthrich K.
        J. Mol. Graphics. 1996; 14: 51-55