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• W2017757947 abstract "Four double mutants in the ε subunit were generated, each containing two cysteines, which, based on the NMR structure of this subunit, should form internal disulfide bonds. Two of these were designed to generate interdomain cross-links that lock the C-terminal α-helical domain against the β-sandwich (εM49C/A126C and εF61C/V130C). The second set should give cross-linking between the two C-terminal α-helices (εA94C/L128C and εA101C/L121C). All four mutants cross-linked with 90–100% efficiency upon CuCl2 treatment in isolated Escherichia coli ATP synthase. This shows that the structure obtained for isolated ε is essentially the same as in the assembled complex.Functional studies revealed increased ATP hydrolysis after cross-linking between the two domains of the subunit but not after cross-linking between the C-terminal α-helices. None of the cross-links had any effect on proton pumping-coupled ATP hydrolysis, on DCCD sensitivity of this activity, or on ATP synthesis rates. Therefore, big conformational changes within ε can be ruled out as a part of the enzyme function. Protease digestion studies, however, showed that subtle changes do occur, since the ε subunit could be locked in an ADP or 5′-adenylyl-β,γ-imidodiphosphate conformation by the cross-linking with resulting differences in cleavage rates. Four double mutants in the ε subunit were generated, each containing two cysteines, which, based on the NMR structure of this subunit, should form internal disulfide bonds. Two of these were designed to generate interdomain cross-links that lock the C-terminal α-helical domain against the β-sandwich (εM49C/A126C and εF61C/V130C). The second set should give cross-linking between the two C-terminal α-helices (εA94C/L128C and εA101C/L121C). All four mutants cross-linked with 90–100% efficiency upon CuCl2 treatment in isolated Escherichia coli ATP synthase. This shows that the structure obtained for isolated ε is essentially the same as in the assembled complex. Functional studies revealed increased ATP hydrolysis after cross-linking between the two domains of the subunit but not after cross-linking between the C-terminal α-helices. None of the cross-links had any effect on proton pumping-coupled ATP hydrolysis, on DCCD sensitivity of this activity, or on ATP synthesis rates. Therefore, big conformational changes within ε can be ruled out as a part of the enzyme function. Protease digestion studies, however, showed that subtle changes do occur, since the ε subunit could be locked in an ADP or 5′-adenylyl-β,γ-imidodiphosphate conformation by the cross-linking with resulting differences in cleavage rates. A proton translocating F1F0 type ATP synthase can be found in the periplasmic membrane of bacteria, the thylakoid membrane of chloroplasts, and the cristae membranes of mitochondria. This enzyme can use a proton gradient to synthesize ATP, a process that is reversible in bacteria where the hydrolysis of ATP is used to generate a proton motive force for substrate and ion transport (1Boyer P.D. Annu. Rev. Biochem. 1997; 66: 717-749Crossref PubMed Scopus (1553) Google Scholar, 2Senior A.E. Annu. Rev. Biophys. Biophys. Chem. 1990; 19: 7-41Crossref PubMed Scopus (328) Google Scholar). The best characterized F1F0 type ATP synthase is from Escherichia coli. It is composed of two parts: a membrane-embedded F0 part containing three different subunits (a, b2, c12) (3Foster D.L. Fillingame R.H. J. Biol. Chem. 1982; 257: 2009-2015Abstract Full Text PDF PubMed Google Scholar, 4Jones P.C. Fillingame R.H. J. Biol. Chem. 1998; 273: 29701-29705Abstract Full Text Full Text PDF PubMed Scopus (111) Google Scholar, 5Schneider E. Altendorf K. Microbiol. Rev. 1987; 51: 477-497Crossref PubMed Google Scholar) and a water-soluble ECF1 1The abbreviations used are:ECF1F0ECF1, and ECF0,E. coli ATP synthase and F1 and F0portions, respectivelyAMP-PNP5′-adenylyl-β,γ-imidodiphosphateDCCDdicyclohexylcarbodiimideDTTdithiothreitolE5Meosin 5-maleimideMOPS4-morpholinepropanesulfonic acidNEMN-ethylmaleimideACMA9-amino-6-chloro-2- methoxyacridine 1The abbreviations used are:ECF1F0ECF1, and ECF0,E. coli ATP synthase and F1 and F0portions, respectivelyAMP-PNP5′-adenylyl-β,γ-imidodiphosphateDCCDdicyclohexylcarbodiimideDTTdithiothreitolE5Meosin 5-maleimideMOPS4-morpholinepropanesulfonic acidNEMN-ethylmaleimideACMA9-amino-6-chloro-2- methoxyacridine part with five different subunits (α3, β3, γ, δ, ε). The F0 portion forms the proton channel (6Elston T. Wang H. Oster G. Nature. 1998; 391: 510-514Crossref PubMed Scopus (438) Google Scholar, 7Dimroth P. Wang H. Grabe M. Oster G. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 4924-4929Crossref PubMed Scopus (132) Google Scholar). The F1 portion contains the three catalytic sites, each predominantly on a β subunit, and located at the interfaces with the α subunits. Recent electron microscopy studies show that the F1 and F0 parts are connected by two stalks (8Wilkens S. Capaldi R.A. Biochim. Biophys. Acta. 1998; 1365: 93-97Crossref PubMed Scopus (75) Google Scholar,9Böttcher B. Schwarz L. Gräber P.J. J. Mol. Biol. 1998; 281: 757-762Crossref PubMed Scopus (87) Google Scholar), a rotating central stalk involving the γ and ε subunits (10Tang C. Capaldi R.A. J. Biol. Chem. 1996; 271: 3018-3024Abstract Full Text Full Text PDF PubMed Scopus (79) Google Scholar,11Aggeler R. Ogilvie I. Capaldi R.A. J. Biol. Chem. 1997; 272: 19621-19624Abstract Full Text Full Text PDF PubMed Scopus (104) Google Scholar) and an outer stalk of the b and δ subunits (12Ogilvie I. Aggeler R. Capaldi R.A. J. Biol. Chem. 1997; 272: 16652-16656Abstract Full Text Full Text PDF PubMed Scopus (144) Google Scholar, 13Rodgers A.J. Capaldi R.A. J. Biol. Chem. 1998; 273: 29406-29410Abstract Full Text Full Text PDF PubMed Scopus (76) Google Scholar). This more peripheral stalk is thought to act as a stator that holds the α-β hexagon in position, while the central stalk is rotating inside the α-β subunits (14Sabbert D. Engelbrecht S. Junge W. Nature. 1996; 381: 623-625Crossref PubMed Scopus (459) Google Scholar, 15Noji H. Yasuda R. Yoshida M. Kinosita Jr., K. Nature. 1997; 386: 299-302Crossref PubMed Scopus (1924) Google Scholar, 16Bulygin V.V. Duncan T.M. Cross R.L. J. Biol. Chem. 1998; 273: 31765-31969Abstract Full Text Full Text PDF PubMed Scopus (50) Google Scholar). ECF1, and ECF0,E. coli ATP synthase and F1 and F0portions, respectively 5′-adenylyl-β,γ-imidodiphosphate dicyclohexylcarbodiimide dithiothreitol eosin 5-maleimide 4-morpholinepropanesulfonic acid N-ethylmaleimide 9-amino-6-chloro-2- methoxyacridine ECF1, and ECF0,E. coli ATP synthase and F1 and F0portions, respectively 5′-adenylyl-β,γ-imidodiphosphate dicyclohexylcarbodiimide dithiothreitol eosin 5-maleimide 4-morpholinepropanesulfonic acid N-ethylmaleimide 9-amino-6-chloro-2- methoxyacridine The structures of the α and β subunits are known in detail from x-ray crystallography of both bovine heart and rat liver mitochondrial F1 (17Abrahams J.P. Leslie A.G. Lutter R. Walker J.E. Nature. 1994; 370: 621-628Crossref PubMed Scopus (2718) Google Scholar, 18Bianchet M.A. Hullihen J. Pedersen P.L. Amzel L.M. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 11065-11070Crossref PubMed Scopus (224) Google Scholar). Part of the γ subunit was resolved in the bovine heart F1, but neither of these structures provided information about the ε subunit (δ subunit of mammalian F1). However, the structure of the ε subunit when isolated from the E. coli enzyme has recently been solved by both NMR and x-ray crystallography (19Wilkens S. Dahlquist F.W. McIntosh L.P. Donaldson L.W. Capaldi R.A. Nat. Struct. Biol. 1995; 2: 961-967Crossref PubMed Scopus (155) Google Scholar, 20Uhlin U. Cox G.B. Guss J.M. Structure. 1997; 5: 1219-1230Abstract Full Text Full Text PDF PubMed Scopus (154) Google Scholar, 21Wilkens S. Capaldi R.A. J. Biol. Chem. 1998; 273: 26645-26651Abstract Full Text Full Text PDF PubMed Scopus (93) Google Scholar). This subunit is a two-domain protein with an N-terminal part arranged in a β-sandwich structure and the C-terminal portion in a helix-loop-helix motif. To understand how the ε subunit functions in the intact ECF1F0, it is necessary to know whether the structure of ε is the same in the assembled complex as when isolated. To examine this question, we have created four different mutants, each containing two cysteines, that should form disulfide bridges if the structure of the ε subunit is the same as in solution. The effects of generating disulfide bridges within the ε subunit on the functioning of ECF1F0 have also been examined. Site-directed mutagenesis was carried out according to Kunkel et al. (22Kunkel T.A. Roberts J.D. Zakour M.A. Methods. Enzymol. 1987; 154: 367-382Crossref PubMed Scopus (4540) Google Scholar) using CJ236 (New England Biolabs). For routine cloning procedures, the strain XLI-Blue (Stratagene) was used (23Maniatis T. Fritsch E.F. Sambrook J. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1982Google Scholar, 24Davis L.G. Dibner M.D. Battey J.F. Basic Methods in Molecular Biology. Elsevier, New York1986Google Scholar). The double mutations in ε were created using two oligonucleotides (see Table I) with M13mp18 that contained the wild-type ε gene. Screening for mutants was done using the appropriate restriction enzymes. Subcloning of the mutated ε gene into pRA100 was performed as described by Aggeler et al. (25Aggeler R. Chicas-Cruz K. Cai S.X. Keana J.F.W. Capaldi R.A. Biochemistry. 1992; 31: 2956-2961Crossref PubMed Scopus (95) Google Scholar).Table IList of oligonucleotides used for site-directed mutagenesisMutantsOligonucleotideRestriction enzymeM49CC ATT AAG CCT GGT TGTHinfI (10214Δ)ATT CGC ATC GTGA126CCTG GCC AAA GCG ATCTGTBsiEI (10441Δ)CAG CTG CGC GTT ATC GF61CG CAC GGT CAC GAA GAGTGC ATC TAT CTG TCT GGCSfaNI (10257+)V130CGCG CAG CTG CGC TGC ATCGAG TTG ACC AAA AAA GCGSfaNI (10465+)A94CGAC GAA GCG CGA TGTNcoI (10348Δ)ATG GAA GCG AAA CGL128CGCG ATC GCG CAG TGT CGCPvuII (10448Δ)GTT ATC GAGA101CGCG AAA CGT AAG TGCEarI (10366Δ)GAG GAG CAC ATT AGCL121CGCG TCT GCG GAA TGT GCCEaeI (10429Δ)AAA GCG ATC GChanged bases are underlined; affected restriction sites are listed accordingly (Δ, missing site; +, additional site); numbering according to Walker et al. (26Walker J.E. Saraste M. Gay N.J. Biochim. Biophys. Acta. 1984; 768: 164-200Crossref PubMed Scopus (370) Google Scholar). Open table in a new tab Changed bases are underlined; affected restriction sites are listed accordingly (Δ, missing site; +, additional site); numbering according to Walker et al. (26Walker J.E. Saraste M. Gay N.J. Biochim. Biophys. Acta. 1984; 768: 164-200Crossref PubMed Scopus (370) Google Scholar). Restriction enzymes used were purchased from Roche Molecular Biochemicals or New England Biolabs. Reconstituted vesicles were washed in DTT-free buffer A (25 mm MOPS/HCl, 5 mmMgCl2, 10% glycerol, pH 7.0) before inducing cross-linking with varying concentrations of CuCl2 (1 h, room temperature). The reaction was stopped with 10 mm EDTA (5 min) before 25 μm E5M was added (5 min, room temperature, in the dark). The reaction was quenched by saturating the unreacted cysteines with 10 mm NEM (5 min, room temperature). As a zero control, washed, reconstituted vesicles were reacted with E5M directly. A second control was incubated with 10 mm DTT (1 h, room temperature) and then quenched by adding 20 mm NEM. To all samples, DTT-free dissociation buffer was added and then subjected to 10–20% SDS-polyacrylamide gel electrophoresis. The same procedure was applied to inner membranes where necessary. Reconstituted ECF1F0 was washed and CuCl2-treated as described above. Then the uncross-linked cysteines were reacted with NEM (10 mm) for 10 min at room temperature, and the volume was raised to 1 ml with buffer A, followed by centrifugation at 60,000 rpm, 20 min, 4 °C in a Beckmann TLA100.2 rotor. This procedure was repeated once before adding 5 mm DTT (20 min, room temperature) to open the disulfide bridges in the ε subunit. After repeating the washing step twice, free cysteines were reacted with 25 μm E5M for 10 min at room temperature in the dark, and the reaction was quenched with 10 mm NEM. Noncross-linked ECF1F0 was used as a control. All samples were applied on a 10–20% polyacrylamide gel as before. Preweighed trypsin was purchased from Roche Molecular Biochemicals. Reconstituted vesicles were washed in 25 mm Tris/HCl, 5 mm MgCl2, 10% glycerol, pH 7.5, to remove DTT and resuspended in the same buffer at 1 mg/ml. Samples were preincubated for 5 min at room temperature in the presence or absence of 5 mm AMP-PNP. Cross-linking was done for 1 h at room temperature in the presence of 75 μmCuCl2. The reaction was stopped with 7 mm EDTA, and Mg2+ AMP-PNP was added to a final concentration of 5 mm (5 min, room temperature) before the trypsinization was started (ECF1F0:trypsin, 50:1). The cleavage was stopped at different times by adding 2 μmphenylmethanesulfonyl fluoride. The same procedure was applied to noncross-linked ECF1F0. Samples were applied on a 10–20% polyacrylamide gel either in the presence or absence of DTT. 50 μl (1 mg/ml) of inner membranes were diluted 10-fold in buffer B (10 mm Hepes, 5 mmMgCl2, 100 mm KCl, pH 7.0), and then 1 μl of valinomycin (1.8 mm), 5 μl of ACMA (0.1 mm), 5 μl of NADH2 (50 mm), and 5 μl of KCN (200 mm) were added. The fluorescence at 480 nm was measured for a short time before adding 5 μl of ATP (200 mm). After the signal reached a plateau, 1 μl of nigericin (1.8 mm) was added to uncouple the system. The excitation wavelength was 410 nm. For ATP-dependent proton pumping measurements, an SLM 8000 fluorometer was used. Inner membranes were washed twice with 25 mm Tris/HCl, 10% glycerol, 5 mmMgCl2, pH 7.5, diluted to 1 mg/ml, and reacted with 75 μm CuCl2 for 1 h at room temperature if necessary. 200 μl of Tris buffer were added to 50 μl inner membranes (50 μg) followed by 250 μl of reaction buffer (100 mm Tris/HCl, 10 mm ADP, 5 mmMgCl2, 10 mm K2HPO4, pH 7.5). 100-μl samples were immediately quenched with 30 μl of 0.5m trichloroacetic acid on ice to measure the amount of endogenous ATP. To the residual 400-μl sample, 50 μmNADH (4 μl, 0.5 m stock) was added and incubated in a 37 °C water bath. The reaction was stopped after 5 min by adding 130 μl, 0.5 m trichloroacetic acid on ice. All samples including the background controls were diluted 25-fold with a buffer containing 0.1 m Tris acetate, 2 mm EDTA, pH 7.7, and measured. The amount of ATP was determined for 100 μl of diluted sample, adding 50 μl of a 1:2 diluted luciferin/luciferase ATP assay mix (Sigma) and 150 μl of Tris acetate buffer. The emitted light was detected using a chemiluminometer after standardizing with preweighed ATP (also purchased from Sigma). Inner membranes were isolated from the strain RA1 (11Aggeler R. Ogilvie I. Capaldi R.A. J. Biol. Chem. 1997; 272: 19621-19624Abstract Full Text Full Text PDF PubMed Scopus (104) Google Scholar) according to the procedure of Foster and Fillingame (27Foster D.L. Fillingame R.H. J. Biol. Chem. 1979; 254: 8230-8236Abstract Full Text PDF PubMed Google Scholar). For the purification of ECF1F0, a method described by Foster and Fillingame (27Foster D.L. Fillingame R.H. J. Biol. Chem. 1979; 254: 8230-8236Abstract Full Text PDF PubMed Google Scholar) and modified by Aggeleret al. (28Aggeler R. Zhang Y.Z. Capaldi R.A. Biochemistry. 1987; 26: 7107-7113Crossref PubMed Scopus (27) Google Scholar) was used. ATP hydrolysis was measured using a regenerating system (29Lötscher H.-R. deJong C. Capaldi R.A. Biochemistry. 1984; 23: 4140-4143Crossref PubMed Scopus (99) Google Scholar). Protein concentrations were determined using the test of Sedmak and Grossberg (30Sedmak J.J. Grossberg S.E. Anal. Biochem. 1977; 79: 544-552Crossref PubMed Scopus (2471) Google Scholar). Reconstitution of ECF1F0 into Egg-PC vesicles was performed as described (31Aggeler R. Haughton M.A. Capaldi R.A. J. Biol. Chem. 1995; 270: 9185-9191Abstract Full Text Full Text PDF PubMed Scopus (121) Google Scholar). DTT was removed by centrifugation in a Beckman TLA100.2 rotor, for 20 min, 60,000 rpm, at 4 °C. The pellet was resuspended in DTT-free buffer and centrifuged again. The step was repeated and the pellets resuspended in DTT-free buffer at a final concentration of 0.5–1 mg/ml. Samples were analyzed on 10–20% SDS-polyacrylamide gel electrophoresis (32Laemmli U.K. Nature. 1970; 227: 680-685Crossref PubMed Scopus (205523) Google Scholar) and stained with Coomassie Brilliant Blue R (33Downer N.W. Robinson N.C. Capaldi R.A. Biochemistry. 1976; 15: 2930-2936Crossref PubMed Scopus (176) Google Scholar). ACMA and E5M were purchased from Molecular Probes, Inc. (Eugene, OR). All other chemicals were of the highest purity available. Four different double mutants were used in this study, two with cysteine residues introduced at sites that are at the interface between the two domains (M49C/A126C or F61C/V130C, respectively) and two that place the two cysteines in close proximity at the interacting faces of the C-terminal α-helices (A94C/L128C or A101C/L121C). Fig.1 shows the position of these substitutions in the ε solution structure. With ECF1F0 purified from each mutant, disulfide bond formation was readily obtained in essentially 100% yield when high enough levels of Cu2+ were used. The SDS-polyacrylamide gel in Fig.2 A shows the effect of internal cross-linking on the migration of ε for the mutant εM49C/A126C. CuCl2 treatment led to a shift to apparent lower molecular weight but, at the same time, to a weaker overall staining of the ε subunit band. This altered staining intensity made it difficult to quantitate cross-link yields. Therefore, the formation of the disulfide bond was also followed by fluorescence after labeling with E5M. Fig. 2, B and C, shows the result of E5M labeling of εM49C/A126C using two different approaches. In one approach, cross-linking was induced by CuCl2, and the amount of ε that had not become cross-linked was determined by modification of remaining exposed cysteines with E5M (Fig. 2 B). As a complementary approach, cross-linked enzyme was reacted with NEM to block free cysteines, and then the disulfide bridge was broken by adding DTT to expose cysteine residues previously cross-linked. These were then modified with E5M (Fig. 2 C). Both approaches gave the same results. High cross-linking yields with the mutant εM49C/A126C required 75 μm CuCl2. Fig. 2,D and E, shows cross-linking between the C-terminal helices in the mutant εA101C/L121C. In this case, uncross-linked ε labeled with E5M gave a diffuse band (D, lane 3), an effect prevented by previous cross-link formation. As with the other mutants, cross-link yields were quantitated by E5M incorporation. Essentially, a 100% yield of cross-linking could be obtained at 10 μmCuCl2. The effects of cross-linking within the ε subunit on the function of the ATP synthase are summarized in Fig.3 A. Those mutants that formed a cross-link between N- and C-terminal domains had ATPase activities similar to wild-type enzyme. As shown for εM49C/A126C in Fig.3 B, ATPase activity increased in proportion to the cross-linking yield, giving a 2.5-fold higher turnover rate. Nevertheless, the cross-linked and activated enzyme retained similar levels of DCCD inhibition to untreated or wild-type enzyme. Moreover, this activation of ATPase was not accompanied by enhanced steady-state levels of ATP-dependent proton pumping when measured by ACMA quenching (Fig. 4) or of ATP synthesis (Fig. 3) when these activities were measured in inner membranes from E. coli. The ATPase activity of the mutant εA94C/L128C was significantly higher than wild-type even before the addition of Cu2+ and essentially the same as that of fully cross-linked enzyme from the mutant εM49C/A126C. Cu2+treatment and the resulting cross-linking of εA94C/L128C in yields greater than 90% failed to increase activity much further. In contrast, the ATPase activity of the mutant εA101C/L121C was normal and not significantly activated by cross-linking. The DCCD sensitivity of both εA94C/L128C and εA101C/L121C was similar to wild-type (Fig.4). For both of these mutants, disulfide bond formation between the two C-terminal α helices had no significant effect on proton pumping or ATP synthesis.Figure 4ATP-dependent ACMA quenching of wild-type (A), εM49C/A126C (B), εA94C/L128C (C), and εA101C/L121C (D) in the presence of 5 mm DTT (solid lines) or 100 μm CuCl2 (dashed lines). All mutants had a yield of at least 90% internal cross-link based on E5M fluorescence.View Large Image Figure ViewerDownload Hi-res image Download (PPT) The nucleotide dependence of cross-linking of the mutant εM49C/A126C is shown in Fig. 5. Disulfide bond formation was most efficient with 5 mm Mg2+ AMP-PNP present. In the presence of Mg2+ATP, which is rapidly hydrolyzed during the experiment, Mg2+ADP, or Mg2+ADP + Pi (5 mm respectively), the yields of cross-linking were 40–50%, compared with 90–100% in AMP-PNP. This can be seen clearly in the Coomassie-stained SDS-polyacrylamide gels (Fig. 5 A). In Mg2+AMP-PNP (lane 2), there is a nearly complete bandshift, whereas in ADP, ADP + Pi, or ATP (lanes 4–6), two bands are clearly visible. These conclusions are confirmed by E5M labeling (Fig. 5 B). Cross-linking of the mutant εF61C/V130C gave the same general patterns as for εM49C/A126C. The nucleotide dependence of cross-linking of mutants with both Cys in the C-terminal part was not examined in any detail. Earlier studies had shown a nucleotide dependence of the cleavage of ε by trypsin in the wild-type ECF1F0 complex (34Mendel-Hartvig J. Capaldi R.A. Biochemistry. 1991; 30: 10987-10991Crossref PubMed Scopus (55) Google Scholar), and this proteolysis was found to activate the enzyme. Cleavage of the ε subunit was followed in the mutant εM49C/A126C in the presence of Mg2+ADP and in Mg2+AMP-PNP. As with wild-type enzyme, cleavage of uncross-linked enzyme was fast in the presence of AMP-PNP but slow in the presence of Mg2+ADP. If both the mutant was cross-linked and protease digestion was conducted in AMP-PNP, the rate of cleavage was fast. However, if cross-linking occurred in the presence of Mg2+ADP, and then the ADP was replaced by AMP-PNP before trypsin treatment, the cleavage was slow (Fig. 6). Clearly, cross-linking in ADP traps the ε subunit in a conformation that is not subsequently modified by the addition of AMP-PNP. The inaccessibility of trypsin to cleavage sites could be a direct result of fixing the two domains of ε together so that the C-terminal α helix cannot unfold. Alternatively, the cross-link could fix a conformation of the whole ECF1F0 where the cleavage sites in ε are sterically shielded by other subunits. The present study offers important insights into the arrangement and functioning of the ε subunit in ECF1F0. Four mutants were generated, by reference to the structure of ε from NMR studies of the isolated polypeptide (21Wilkens S. Capaldi R.A. J. Biol. Chem. 1998; 273: 26645-26651Abstract Full Text Full Text PDF PubMed Scopus (93) Google Scholar). In each of these mutants, the positions of the cysteines are such that they should readily form disulfide bonds if the observed structure is retained in the enzyme complex. Internal cross-links were readily formed in essentially 100% yield between the introduced Cys in all four mutants when ECF1F0 was reacted with Cu2+. Two of the disulfide bridges are between one residue in the N-terminal β-barrel part and the very C-terminal α-helix of the C-terminal domain of the subunit. The other two disulfide bridges are between a residue in each of the C-terminal α-helices. This ready cross-linking would only be expected if the two domains and both C-terminal helices have the same position in ECF1F0 as in isolated ε subunit. Therefore, it can be concluded that the structure of ε in the intact complex is very similar to that already determined for purified ε subunit. The ε subunit, along with γ, is now thought to be a part of the rotor that couples catalytic site events with proton pumping in the complex (11Aggeler R. Ogilvie I. Capaldi R.A. J. Biol. Chem. 1997; 272: 19621-19624Abstract Full Text Full Text PDF PubMed Scopus (104) Google Scholar, 35Schulenberg B. Wellmer F. Lill H. Junge W. Engelbrecht S. Eur. J. Biochem. 1997; 249: 134-141Crossref PubMed Scopus (46) Google Scholar). Nucleotide-dependent changes in interaction of the ε subunit with other subunits in ECF1F0 have been resolved by cross-linking studies (36Aggeler R. Capaldi R.A. J. Biol. Chem. 1996; 271: 13888-13891Abstract Full Text Full Text PDF PubMed Scopus (102) Google Scholar). These involve both domains of the subunit, and it has been suggested that the two domains move independently during functioning of the enzyme (35Schulenberg B. Wellmer F. Lill H. Junge W. Engelbrecht S. Eur. J. Biochem. 1997; 249: 134-141Crossref PubMed Scopus (46) Google Scholar). The observation that the two domains can be tethered together in either of two different places without altering coupling argues against this possibility but does not rule out subtle rotations of one domain relative to the other. The cross-linking studies here rule out that the C-terminal α-helices come apart during functioning (21Wilkens S. Capaldi R.A. J. Biol. Chem. 1998; 273: 26645-26651Abstract Full Text Full Text PDF PubMed Scopus (93) Google Scholar). This is not to say that conformational changes are absent from ε. The nucleotide dependence of cross-linking in mutants forming S–S bridges between the two domains points to some structural changes within ε. This notion is also supported by the observation that rates of cleavage of ε by trypsin are affected by the nucleotide conditions in which cross-linking is conducted prior to the proteolysis step. When cross-linked in ADP the ε subunit was locked in a conformation characterized by very slow proteolysis, a conformation that was not altered by the addition of AMP-PNP before trypsin treatment. In the presence of AMP-PNP, proteolysis of the ε subunit is normally rapid. The effect of cross-links within the ε subunit on functioning of ECF1F0 adds to our understanding of the role(s) of the two domains of this subunit. Cross-linking to fix the N- and C-terminal domains increases ATP hydrolysis by the enzyme complex up to 2.5- fold. The introduction of Cys for Ala94 plus Leu128 also increases ATPase activity more than 2-fold, an activation that is not greatly enhanced by disulfide bond formation between the introduced Cys residues. The activation of ATP hydrolysis in these mutants is without effect on coupling because, in each case, the steady-state levels of proton pumping, as well as ATP synthesis rates, are unaltered. Therefore, our results support and extend previous studies that indicate that the ε subunit acts to regulate ATPase rates in the intact ATP synthase, i.e. that it is an inhibitor of this function in the intact complex (34Mendel-Hartvig J. Capaldi R.A. Biochemistry. 1991; 30: 10987-10991Crossref PubMed Scopus (55) Google Scholar, 38Richter M.L. Snyder B. McCarty R.E. Hammes G.G. Biochemistry. 1985; 24: 5755-5763Crossref PubMed Scopus (85) Google Scholar) (for the opposite view, see Refs. 39Sternweis P.C. Smith J.B. Biochemistry. 1980; 19: 526-531Crossref PubMed Scopus (106) Google Scholar and 40Ketchum C.J. Nakamoto R.K. J. Biol. Chem. 1998; 273: 22292-22297Abstract Full Text Full Text PDF PubMed Scopus (24) Google Scholar). This inhibitor function is probably provided by the N-terminal helical domain. Protease digestion of this domain by trypsin in ECF1 (41Mendel-Hartvig J. Capaldi R.A. Biochemistry. 1991; 30: 1278-1284Crossref PubMed Scopus (71) Google Scholar) alters the inhibition by the ε subunit, while proteolysis and genetic deletion of part or all of this domain has no effect on coupling function (42Cruz J.A. Harfe B. Radkowski C.A. Dann M.S. McCarty R.E. Plant Physiol. 1995; 109: 1379-1388Crossref PubMed Scopus (50) Google Scholar, 43Kuki M. Noumi T. Maeda M. Amemura A. Futai M. J. Biol. Chem. 1988; 263: 17437-17442Abstract Full Text PDF PubMed Google Scholar, 44Xiong H. Zhang D. Vik S.B. Biochemistry. 1998; 37: 16423-16429Crossref PubMed Scopus (42) Google Scholar). Our previous studies have shown that the C-terminal domain lies under the α3β3 hexagon in the enzyme complex, where it associates with two α−β pairs (21Wilkens S. Capaldi R.A. J. Biol. Chem. 1998; 273: 26645-26651Abstract Full Text Full Text PDF PubMed Scopus (93) Google Scholar, 36Aggeler R. Capaldi R.A. J. Biol. Chem. 1996; 271: 13888-13891Abstract Full Text Full Text PDF PubMed Scopus (102) Google Scholar). This domain could regulate ATP hydrolysis by controlling catalytic site cooperativity. The N-terminal β barrel domain binds to the γ subunit (10Tang C. Capaldi R.A. J. Biol. Chem. 1996; 271: 3018-3024Abstract Full Text Full Text PDF PubMed Scopus (79) Google Scholar) and the c subunit ring (37Zhang Y. Fillingame R.H. J. Biol. Chem. 1995; 270: 24609-24614Abstract Full Text Full Text PDF PubMed Scopus (102) Google Scholar), as would be expected of a key component of the coupling mechanism. We thank Dr. Robert Aggeler for many helpful discussions and for help with some of the mutants used here. Also, Kathy Chicas-Cruz and Beth Monika are thanked for help in making enzyme preparations." @default.
• W2017757947 created "2016-06-24" @default.
• W2017757947 creator A5057987370 @default.
• W2017757947 creator A5080317638 @default.
• W2017757947 date "1999-10-01" @default.
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