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• W2154767191 abstract "The binding change model for the F1-ATPase predicts that its rotation is intimately correlated with the changes in the affinities of the three catalytic sites for nucleotides. If so, subtle differences in the nucleotide structure may have pronounced effects on rotation. Here we show by single-molecule imaging that purine nucleotides ATP, GTP, and ITP support rotation but pyrimidine nucleotides UTP and CTP do not, suggesting that the extra ring in purine is indispensable for proper operation of this molecular motor. Although the three purine nucleotides were bound to the enzyme at different rates, all showed similar rotational characteristics: counterclockwise rotation, 120° steps each driven by hydrolysis of one nucleotide molecule, occasional back steps, rotary torque of ∼40 piconewtons (pN)·nm, and mechanical work done in a step of ∼80 pN·nm. These latter characteristics are likely to be determined by the rotational mechanism built in the protein structure, which purine nucleotides can energize. With ATP and GTP, rotation was observed even when the free energy of hydrolysis was −80 pN·nm/molecule, indicating ∼100% efficiency. Reconstituted FoF1-ATPase actively translocated protons by hydrolyzing ATP, GTP, and ITP, but CTP and UTP were not even hydrolyzed. Isolated F1 very slowly hydrolyzed UTP (but not CTP), suggesting possible uncoupling from rotation. The binding change model for the F1-ATPase predicts that its rotation is intimately correlated with the changes in the affinities of the three catalytic sites for nucleotides. If so, subtle differences in the nucleotide structure may have pronounced effects on rotation. Here we show by single-molecule imaging that purine nucleotides ATP, GTP, and ITP support rotation but pyrimidine nucleotides UTP and CTP do not, suggesting that the extra ring in purine is indispensable for proper operation of this molecular motor. Although the three purine nucleotides were bound to the enzyme at different rates, all showed similar rotational characteristics: counterclockwise rotation, 120° steps each driven by hydrolysis of one nucleotide molecule, occasional back steps, rotary torque of ∼40 piconewtons (pN)·nm, and mechanical work done in a step of ∼80 pN·nm. These latter characteristics are likely to be determined by the rotational mechanism built in the protein structure, which purine nucleotides can energize. With ATP and GTP, rotation was observed even when the free energy of hydrolysis was −80 pN·nm/molecule, indicating ∼100% efficiency. Reconstituted FoF1-ATPase actively translocated protons by hydrolyzing ATP, GTP, and ITP, but CTP and UTP were not even hydrolyzed. Isolated F1 very slowly hydrolyzed UTP (but not CTP), suggesting possible uncoupling from rotation. the α3β3γ subcomplex (β-His tag/γS107C) of F1 derived from thermophilicBaccillus strain PS3 nitrilotriacetic acid carbonyl cyanide p-(trifluoromethoxy) phenylhydrazone FoF1-ATPase derived from thermophilic Baccillus strain PS3 4-morpholinepropanesulfonic acid piconewton(s) The FoF1 ATP synthase is an enzyme that synthesizes ATP from ADP and inorganic phosphate (Pi) using proton flow across a membrane (1Boyer P.D. Biochim. Biophys. Acta. 2000; 1458: 252-262Crossref PubMed Scopus (98) Google Scholar). The Fo portion of the enzyme resides in the membrane and mediates proton translocation. The F1 portion, consisting of α3β3γ1δ1ε1subunits, is external to the membrane and catalyzes ATP synthesis. The ATP synthase is a completely reversible molecular machine in that ATP hydrolysis in F1 can produce a reverse flow of protons through Fo. Isolated F1 only catalyzes ATP hydrolysis and hence is called F1-ATPase. Its minimal, stable subcomplex capable of ATP hydrolysis is α3β3γ (2Matsui T. Yoshida M. Biochim. Biophys. Acta. 1995; 1231: 139-146Crossref PubMed Scopus (86) Google Scholar). For the coupling between the proton flow in Fo and the chemical reaction (ATP synthesis/hydrolysis) in F1, Boyer and Kohlbrenner (3Boyer P.D. Kohlbrenner W.E. Selman B.R. Selman-Reimer S. Energy Coupling in Photosynthesis. Elsevier, Amsterdam1981: 231-240Google Scholar) and Oosawa and Hayashi (4Oosawa F. Hayashi S. Adv. Biophys. 1986; 22: 151-183Crossref PubMed Scopus (98) Google Scholar) independently suggested a rotational catalysis model. The essence is that Fo is a rotary motor (or turbine) driven by the proton flow, that F1 is another rotary motor driven by ATP hydrolysis, and that the two motors have a common rotary shaft, yet their genuine rotary directions are opposite to each other's. Rotation of the shaft in Fo's genuine direction, as occurs in cells, results in the reverse rotation of the F1 motor and thus in ATP synthesis in F1. When F1 gains control and hydrolyzes ATP, protons are pumped through Fo in the reverse direction. A crystal structure of F1 (5Abrahams J.P. Leslie A.G. Lutter R. Walker J.E. Nature. 1994; 370: 621-628Crossref PubMed Scopus (2754) Google Scholar) strongly supported the rotation model and has inspired many experiments, which, together, have proved that the γ subunit is (part of) the common rotor shaft and that α3β3 subunits, which surrounded γ in the crystal, are the stator in the F1 motor (6Aggeler R. Haughton M.A. Capaldi R.A. J. Biol. Chem. 1995; 270: 9185-9191Crossref PubMed Scopus (121) Google Scholar, 7Duncan T.M. Bulygin V.V. Zhou Y. Hutcheon M.L. Cross R.L. Proc. Natl. Acad. Sci. U. S. A. 1995; 92: 10964-10968Crossref PubMed Scopus (459) Google Scholar, 8Sabbert D. Engelbrecht S. Junge W. Nature. 1996; 381: 623-625Crossref PubMed Scopus (464) Google Scholar). Single-molecule imaging of F1, in particular, has revealed that the γ subunit rotates in a unique direction consistent with the crystal structure, that γ makes discrete 120° steps, and that the energy conversion efficiency of the F1 motor driven by ATP hydrolysis can reach ∼100% (9Noji H. Yasuda R. Yoshida M. Kinosita Jr., K. Nature. 1997; 386: 299-302Crossref PubMed Scopus (1966) Google Scholar, 10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar). The precise mechanism of rotation, however, is not yet clear. In Boyer's model, binding changes play the major role (1Boyer P.D. Biochim. Biophys. Acta. 2000; 1458: 252-262Crossref PubMed Scopus (98) Google Scholar). On the three β subunits, each of which hosts a catalytic site, ATP and its hydrolysis products, ADP and Pi, are in equilibrium. Thus, in the absence of an external energy supply, the change in free energy associated with ATP hydrolysis should manifest as the higher affinity for ATP than for ADP and Pi. During ATP synthesis, the mechanical energy supplied by the γ rotation driven by F0somehow decreases the affinity for ATP (and probably increases the affinity for ADP and Pi), resulting in the appearance of ATP in the medium. In the reverse reaction of ATP hydrolysis, the free energy difference between the stronger ATP binding and weaker ADP/Pi binding drives the rotation of γ. Boyer proposes that these binding changes occur sequentially on the three catalytic sites, synchronously with the γ rotation. In the crystal structure of F1 (5Abrahams J.P. Leslie A.G. Lutter R. Walker J.E. Nature. 1994; 370: 621-628Crossref PubMed Scopus (2754) Google Scholar), the three β subunits carried a different nucleotide, an analog of ATP, ADP, and none, in support of the sequential binding change mechanism. If binding and release, rather than the cleavage of the terminal phosphate, are indeed the source of mechanical torque production in the F1 motor, the rotational characteristics are expected to differ among different NTPs. The free energy gain in overall hydrolysis is similar among NTPs, but the operation of the F1 motor, which can work at near 100% efficiency, may well depend on delicate energy balances in various steps of hydrolysis. Here we show that purine but not pyrimidine nucleotides support F1 rotation, suggesting that the interaction between the catalytic site and the additional ring in purines is critical to the proper operation of this molecular machine. Nucleotides (GTP, ITP, UTP, and CTP) were purchased from Sigma. ATP and other enzymes were from Roche Molecular Biochemicals. The purity of each nucleotide was assessed on an anion exchange column. Nucleotides were applied on DEAE 2SW (Tosoh, Japan) equilibrated with 50 mm sodium phosphate (pH 7.0) and eluted with an isocratic flow of the same buffer. Because DEAE 2SW has higher affinity for compounds with more negative charges, nucleotides are eluted in the order of nucleoside mono-, di-, and triphosphate. The elution profiles for UTP are shown in Fig.1, a and b. Besides the UDP peak at 13 ml (identical with the peak for commercial UDP), several contaminant peaks (C1–C5) were resolved. The void peak at 4 ml is unlikely to be a nucleotide(s), because A 260(green) and A 280 (red), within an absorbance peak of a nucleotide, were extremely low compared with A 220 (blue). C1 at 7 ml was probably UMP, because it was eluted earlier, and the relative magnitudes of A 220, A 260and A 280 were the same as those for UTP and UDP. C4 was judged as contaminating GTP, because its elution volume and the relative magnitudes of A 220,A 260, and A 280 all matched with those for GTP (Fig. 1c). The GTP contamination amounted to 0.01% of UTP, as determined from the peak area ofA 260. The other contaminants, C2, C3, and C5, were of unknown origin; their signatures did not match with those of ATP, GTP, ITP, dATP, or their diphosphates. C2, C3, and C5 in UTP amounted to 0.03, 0.01, and 0.39%, respectively. Although the levels of contaminants were low, they may nevertheless affect the rotation assay, because the affinity of F1-ATPase for UTP is extremely low (see “Results”). We therefore used the fraction at the UTP peak (23 ml) for the rotation assay. CTP was also found to be contaminated by unknown compounds (1.5%); thus, its peak fraction was used for the rotation assay. ATP, GTP, and ITP contained less than 5% contaminants, which were mostly their hydrolysis products; these were used without purification. The α3β3γ subcomplex (β-His tag/γS107C) of F1 derived from thermophilicBaccillus strain PS3 (TF1)1 was expressed in Escherichia coli as described previously (9Noji H. Yasuda R. Yoshida M. Kinosita Jr., K. Nature. 1997; 386: 299-302Crossref PubMed Scopus (1966) Google Scholar) and purified as follows. The cell lysate containing the enzyme was applied on a Ni2+-NTA Superflow column (Qiagen) equilibrated with 50 mm imidazole (pH 7.0) and 100 mm NaCl. The column was washed with 100 mmimidazole (pH 7.0) and 100 mm NaCl, and then the enzyme was eluted with 500 mm imidazole (pH 7.0) and 100 mm NaCl. Ammonium sulfate was added to the fraction containing the enzyme to a final concentration of 10% saturation and the sample was applied to a butyl-Toyopearl column (Tosoh, Japan) equilibrated with 500 mm imidazole (pH 7.0), 100 mm NaCl, and 10% saturated ammonium sulfate. The column was washed with 10 column volumes of a solution containing 100 mm sodium phosphate (pH 7.0), 2 mm EDTA, and 10% saturated ammonium sulfate to remove endogenously bound nucleotides. The enzyme was eluted with 50 mm Tris-Cl (pH 8.0) and 2 mm EDTA and stored as precipitant in 70% saturated ammonium sulfate containing 2 mm dithiothreitol. Before use, the enzyme was dissolved in 100 mm sodium phosphate (pH 7.0) and 2 mm EDTA and passed through a size exclusion column (Superdex 200 HR 10/30; Amersham Pharmacia Biotech) equilibrated with 100 mm sodium phosphate (pH 7.0) and 2 mm EDTA. The amount of nucleotides remaining on the enzyme was determined as described previously (11Tsunoda S.P. Muneyuki E. Amano T. Yoshida M. Noji H. J. Biol. Chem. 1999; 274: 5701-5706Abstract Full Text Full Text PDF PubMed Scopus (40) Google Scholar). Samples with less than 0.1 mol of nucleotide/mol of enzyme were used for the measurement of hydrolysis activity. Whole TFoF1 complex was reconstituted from the mutant TF1 (β-His tag/γS107C) and authentic Fo from the thermophile and was incorporated into liposomes as described previously (12Bald D. Amano T. Muneyuki E. Pitard B. Rigaud J.L. Kruip J. Hisabori T. Yoshida M. Shibata M. J. Biol. Chem. 1998; 273: 865-870Abstract Full Text Full Text PDF PubMed Scopus (61) Google Scholar). ATP, GTP, ITP, and UTP hydrolysis activities of α3β3γ were measured at 23 °C in a medium containing 50 mm MOPS-KOH (pH 7.0), 50 mm KCl, 2 mm MgCl2, and an ATP (or GTP, ITP, or UTP)-regenerating system consisting of 2.5 mmphosphoenolpyruvate, 100 μg/ml lactate dehydrogenase, 0.2 mm NADH, and pyruvate kinase at 200 μg/ml for ATP hydrolysis, 500 μg/ml for GTP and ITP hydrolysis, or 750 μg/ml for UTP hydrolysis. The reaction was initiated by the addition of the enzyme to 1.3 ml of assay mixture, and hydrolysis was monitored as NADH oxidation determined from the absorbance decrease at 340 nm. The CTP hydrolysis activity was estimated from Pi released in the same medium without a regenerating system during a 60-min incubation at 23 °C. Hydrolysis by the reconstituted TFoF1was also measured as Pi released in 50 mmMOPS-KOH (pH 7.0), 25 mm NaSO4, 25 mm K2SO4, 4 mmMgCl2, and 2 mm indicated nucleotide during a 20-min incubation at 23 °C. Streptavidin-conjugated TF1 for the rotation assay was prepared as described previously (9Noji H. Yasuda R. Yoshida M. Kinosita Jr., K. Nature. 1997; 386: 299-302Crossref PubMed Scopus (1966) Google Scholar). Rotation in the presence of ATP, GTP, or ITP was observed in the TF1adsorbed on a Ni2+-NTA-modified polystyrene bead by attaching an actin filament to the γ subunit (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar). The assay mixture contained 50 mm MOPS-KOH, (pH 7.0), 50 mm KCl, 2 mm MgCl2, 10 mg/ml bovine serum albumin, and an indicated Mg-nucleotide. Fluorescent actin filaments at the bottom of a flow chamber were observed with an inverted epifluorescence microscope (IX70; Olympus). Assays at less than 10 μm GTP or ITP were made on the TF1 directly attached to coverslips to which Ni2+-NTA had been bound covalently (13Adachi K. Yasuda R. Noji H. Itoh H. Harada Y. Yoshida M. Kinosita Jr., K. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 7243-7247Crossref PubMed Scopus (186) Google Scholar); the probability of finding a rotating actin filament was higher than on the Ni2+-NTA-modified polystyrene bead. Images were taken with an intensified CCD camera (ICCD-350F; VideoScope) and recorded on 8-mm videotapes. Rotation by UTP or CTP was observed through an aggregate of biotinylated polystyrene beads with a diameter of 440 nm, which was attached to γ (14Yasuda R. Noji H. Yoshida M. Kinosita Jr., K. Itoh H. Nature. 2001; 410: 898-904Crossref PubMed Scopus (711) Google Scholar). Images of bead aggregates were obtained in the bright field in the absence of the oxygen scavenger system and recorded with a CCD camera (CCD-300-RC; Dage-MTI). Possible nucleotide contamination from actin filaments and the oxygen scavenger system was thus excluded. Nucleotide-driven proton-translocation into TFoF1 liposomes was detected as the decrease in the pyranine fluorescence in an FP 777 fluorometer (JASCO, Japan) at 23 °C. TFoF1liposomes (about 5 μg of TFoF1) containing pyranine inside were preincubated at 23 °C in 1.5 ml of a reaction mixture containing 10 mm MOPS-KOH (pH 7.0), 50 mm KCl, 2 mm nucleotide, 25 mmK2SO4, 25 mmNa2SO4, and 20 mmp-xylene-bispyridinium bromide, which quenched the pyranine fluorescence outside the liposomes. Excitation and emission wavelengths were 460 and 510 nm, respectively. The reaction was started by the addition of 4 mm MgCl2. After recording the fluorescence change, 10 μm FCCP was added to recover the initial fluorescence, which would ensure that the fluorescence decrease was caused by the proton uptake into the liposomes. F1-ATPase is known to be inhibited by Mg-ADP tightly bound to a catalytic site (15Jault J.M. Matsui T. Jault F.M. Kaibara C. Muneyuki E. Yoshida M. Kagawa Y. Allison W.S. Biochemistry. 1995; 34: 16412-16418Crossref PubMed Scopus (67) Google Scholar, 16Matsui T. Muneyuki E. Honda M. Allison W.S. Dou C. Yoshida M. J. Biol. Chem. 1997; 272: 8215-8221Abstract Full Text Full Text PDF PubMed Scopus (102) Google Scholar). TF1 used in the previous study (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar) contained ∼0.3 mol of tightly bound nucleotide/mol of enzyme; thus, the hydrolysis activity might have been underestimated. Here, we employed an improved purification protocol (see “Experimental Procedures”) and obtained a sample containing only 0.084 mol of bound nucleotide (0.037 mol of ATP and 0.047 mol of ADP) per mol of enzyme as determined from the peak heights in the reverse-phase chromatography (Fig.2a). Before use, the protein was further purified by size exclusion chromatography, because TF1 tended to aggregate upon storage. For the final sample, the ratio of A 280 toA 260, a convenient measure of the purity, was greater than 2.0 (Fig. 2b). As expected, the ATP hydrolysis activity of this nucleotide-depleted enzyme was higher than that reported previously (see below). ATP, GTP, ITP, and UTP hydrolysis activities of the nucleotide-depleted TF1 were determined as the rate of oxidation of NADH using the ATP-, GTP-, ITP-, or UTP-regenerating system. Hydrolysis of the purine nucleotides, ATP, GTP, and ITP, gradually decelerated to a steady state as shown in Fig.3, a and b. This turnover-dependent inactivation is attributed to the Mg-ADP (GDP, IDP) inhibition. The hydrolysis rates were therefore determined during the initial 10 s after the injection of the enzyme into the assay mixture (Fig. 3, a and b,insets). The rate of ATP hydrolysis did not obey simple Michaelis-Menten kinetics and was fitted with two sets of K m andV max: V maxa = 87.5 s−1, V maxb= 313 s−1, K m a = 5.25 μm, and K m b = 429 μm (Fig. 3c). The maximum rate of ATP hydrolysis, V maxb, of 313 s−1 is higher than the previous value (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar) of 177 s−1, presumably because of the thorough removal of the bound nucleotide and aggregated enzyme. An additional factor is the increase of the amount of pyruvate kinase, which may limit the overall reaction rate, to 0.2 mg/ml, compared with 0.05 mg/ml in the previous study. The rates for GTP and ITP hydrolysis could be accounted for by simple Michaelis-Menten kinetics;V max and K m were 187 s−1 and 64 μm, respectively, for GTP and 157 s−1 and 202 μm for ITP. In contrast to the purine nucleotides, UTP hydrolysis activity gradually increased with time (Fig. 3, a and b). This cannot be ascribed to the slow response of the UTP-regenerating system, because it could generate UTP from UDP much faster (0.5 s−1) than the acceleration (0.017 s−1 at 1 mm UTP). Slow binding of UTP to noncatalytic sites could explain the acceleration, but evidence is absent. Because the activation took a longer time at lower UTP concentrations, fully activated rate of UTP hydrolysis was estimated from the slope between 150 and 200 s at >300 μm, between 250 and 300 s at 100 μm, and between 1000 and 1200 s at 30 μm. As seen in Fig. 3c, hydrolysis of UTP was much slower than that of the purine nucleotides. To confirm that the measured hydrolysis rate was that of UTP and not of contaminants, we also measured Pi released in the medium. Although the rate of Pi release was around half the hydrolysis rate determined from NADH oxidation, presumably because of the absence of the UTP-regenerating system, TF1 produced Pi equivalent to >40% of UTP in 5 min. Thus, TF1 did hydrolyze UTP, not contaminating other nucleoside-triphosphates, which amounted to <0.5% (see “Experimental Procedures”). CTP, in contrast, was not hydrolyzed; the rate of Pi release at 2 mm CTP measured over 60 min was indistinguishable from the rate without the enzyme (<0.01 s-1). As seen in Fig. 3c, the maximal hydrolysis rates for the four nucleotides, ATP, GTP, ITP, and UTP, do not differ greatly. The major difference among the four is in the apparentK m values, UTP being the highest. In this regard, it is possible that CTP is also hydrolyzed with a similarV max but with a K m far above 10 mm. Another difference is that ATP hydrolysis shows significant deviation from the simple Michaelis-Menten kinetics, which is not apparent for the other nucleotides. This may be correlated with the stronger tendency toward inhibition with ATP than with the other nucleotides (Fig. 3, a and b). The possibility that the other nucleotides also show deviation at much higher concentrations (giving possibly the sameV max) cannot be dismissed. GTP and ITP supported the rotation of the γ subunit in TF1. The rotation, observed through the motion of a fluorescent actin filament attached to the γ subunit, was counterclockwise without exception, as in the case of ATP-driven rotation. Rotation assays at less than 10 μmGTP or 10 μm ITP were made on the enzyme directly attached to a coverslip that had been covalently modified with Ni2+-NTA (13Adachi K. Yasuda R. Noji H. Itoh H. Harada Y. Yoshida M. Kinosita Jr., K. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 7243-7247Crossref PubMed Scopus (186) Google Scholar). Compared with the previous method (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar) of attaching the enzyme on Ni2+-NTA-coated beads, the direct attachment resulted in a higher percentage of finding rotating filaments at low nucleotide concentrations. Because the F1 motor has been shown to be a 120° stepper (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar), we compare 3 times the rotational rate of a short actin filament (0.5–1.3 μm) with the corresponding hydrolysis rate (Fig. 3). At low nucleotide concentrations ([ATP] < 0.1 μm, [GTP] < 1 μm, [ITP] < 10 μm) where nucleotide binding is rate-limiting, the two rates agreed with each other, indicating that three molecules of purine nucleotide, whether ATP, GTP, or ITP, drive a full turn. The agreement also shows that the rotation in the presence of GTP or ITP was not due to possible ATP contamination, which was far below 5%. At higher nucleotide concentrations, the rotational rate saturated around 4 revolutions s−1. This is simply due to the hydrodynamic friction against the rotating actin filament (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar). At low nucleotide concentrations, the rotation was resolved into discrete 120° steps (Fig. 4, a and b). The histograms of dwell times between steps were fitted with a single exponential decay with the rate of 2.9 s−1 for 1 μm GTP and 11.1 s−1 for 10 μm ITP (Fig. 4, c and d), implying that a first order reaction, binding of GTP (ITP), triggers each 120° step. These rate constants agree with the hydrolysis rates (Fig.3c), confirming again that each 120° step is driven by the hydrolysis of one molecule of GTP or ITP. As seen in Fig. 4a, back steps occurred occasionally in GTP-driven rotation. ITP-driven rotation also showed occasional back steps (data not shown). The velocity of back steps was as fast as the forward one and thus is unlikely to be of purely thermal origin (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar). A stochastic mistake in the order of substrate binding or product release among the three catalytic sites could explain the back steps. With commercial UTP at 4 mm, we occasionally observed slow rotation (e.g.1.4 revolutions s−1). We suspected that this might have been due to contaminants, because ATP at a concentration as low as 300 nm, for example, could account for the observed speed. Indeed, even CTP, which the α3β3γ does not hydrolyze, produced rotation at 0.3 revolutions s−1 at 2 mm. We therefore purified UTP and CTP (see “Experimental Procedures”) and examined rotation at 300 μm, the highest concentration available after the column purification. To avoid possible nucleotide contamination from other sources, we attached an aggregate of biotinylated polystyrene beads to the γ subunit in place of an actin filament and observed the beads in bright field in the absence of the oxygen scavenger system. No rotating beads were found in six assays with UTP in which 5543 beads only fluctuated around a fixed point. Observation in each assay continued for more than 30 min, much longer than the time needed to activate UTP hydrolysis at 300 μm (∼5 min). Six assays with CTP also gave negative results. If hydrolysis of each UTP molecule produces a 120° step as with purine nucleotides, the rotational rate is expected to be 4.4 s−1 at 300 μm UTP. We therefore carried out parallel assays with 600 nm ATP where the rotational rate would be 1.5 s−1. 40 rotating beads were found out of 4299 fluctuating beads (five assays). We also made an “unpredicted test” in which we observed beads without knowing the identity of the nucleotide. No rotation was found with 300 μm UTP, and rotating beads were readily found in the presence of 600 nm ATP. We conclude that UTP cannot support rotation of the γ subunit carrying a bead aggregate, at least not at the rate expected from the hydrolysis rate. One possibility is that UTP hydrolysis, unlike the hydrolysis of purine nucleotides, is completely decoupled from γ rotation, even at no load. The other possibility is that UTP, with a different base structure, cannot supply sufficient power to drive the beads through 120°; in this case, the rate of UTP hydrolysis by the loaded TF1 would also be very low. To compare torque and energy conversion efficiency among three purine nucleotides, ATP, GTP, and ITP, that supported γ rotation, the rotational velocities of actin filaments attached to the γ subunit on Ni2+-NTA coated-beads were examined. Fig. 5 shows time courses of rotation for actin filaments with length around 2 μm (Fig.5a) and 3–4 μm (Fig. 5b). The average rotational velocity was determined for each curve over a portion containing at least five consecutive revolutions without noticeably unnatural intermissions (e.g. the last part of thebrown curve in Fig. 5b was omitted). The results are summarized in Fig. 5c. Most data for GTP-driven (green) and ITP-driven rotation (orange) are on or belowthe line representing the constant torque of 40 pN·nm, as is also the case for ATP-driven rotation (blue). We think that higher velocity values are more reliable, because any obstructions against rotation would reduce the velocity, and we conclude that TF1 exerts a constant rotary torque of about 40 pN·nm regardless of the differences among the structures of purine rings. (The average torque calculated over all points in Fig. 5c is 32 ± 11 pN·nm for ATP, 35 ± 11 pN·nm for GTP, and 32 ± 10 pN·nm for ITP; these average values, however, are not meaningful, because the data in Fig. 5c have been selected for high torque values.) The torque of 40 pN·nm times the angular displacement of 2π/3 radians (equal to 120°), ∼80 pN·nm, is the mechanical work done in a 120° step. In the previous study (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar), it was shown that TF1 does this much work even when the free energy available from ATP hydrolysis, ΔG ATP, was reduced to −90 pN·nm/molecule. Here, we examined the torque and work in the presence of 2 mm ATP (or GTP), 10 μm ADP (GDP), and 100 mm inorganic phosphate. Under these conditions, ΔG ATP is calculated as −80 pN·nm/molecule from ΔGATP = ΔG 0 +k B T·ln[ADP]·[Pi]/[ATP], where ΔG0 = −50 pN·nm/molecule (17Stryer L. Biochemistry. 4th Ed. Freeman, New York1995: 443-462Google Scholar),k B T = 4.1 pN·nm/molecule is thermal energy at room temperature, and ΔGGTP should be of a similar value. As seen in Fig. 5c, the experimental points at ΔG ATP = −80 pN·nm/molecule (purple) and ΔG GTP = −80 pN·nm/molecule (brown) are indistinguishable from other points and also fall on or below the line for the constant torque of 40 pN·nm or constant work of 80 pN·nm (the torque averaged over plotted points is 35 ± 10 pN·nm for ΔG ATP = −80 and 32 ± 9 pN·nm for ΔG GTP = −80 pN·nm/molecule) This strengthens our contention that the efficiency of the energy conversion in this motor, from the hydrolysis (of purine nucleotides) into the mechanical work, can reach ∼100%. Nucleotide-driven proton uptake by the reconstituted TFoF1 liposome was measured, to see whether γ rotation is mechanically coupled to the proton pump. Liposomes were incubated with 2 mm nucleotide until the base line became stable, and then 4 mm MgCl2was added to start the reaction (Fig. 6). After 20 min, 10 μm FCCP was added to collapse the pH gradients so as to ensure that the fluorescence decrease indicated the proton uptake. Fig. 6 clearly shows that the purine nucleotides ATP, GTP, and ITP drove the formation of pH gradient but UTP and CTP did not; the latter two gave signals that were indistinguishable from the control in the absence of a nucleotide. The initial rates of proton uptake by GTP and ITP in 1 min after starting the reaction were 30 and 27% of the ATP-driven uptake, respectively. The rate of nucleotide hydrolysis by TFoF1, determined from the amount of inorganic phosphate released in 20 min, was 7.2 ± 1.4 s−1 for ATP, 3.4 ± 0.5 s−1 for GTP, and 2.7 ± 0.5 s−1 for ITP. The three nucleotides show the same order in proton pumping and hydrolysis activities. The agreement suggests that GTP and ITP hydrolysis are coupled to proton pumping as efficiently as with ATP. Pyridine nucleotides were not good substrates for hydrolysis by TFoF1; the hydrolysis activity was not detected by the phosphate measurement (<0.12 s−1 for 2 mm CTP and <0.09 s−1 for 2 mm UTP). In this paper, we have shown that the purine ring of a nucleotide is indispensable for γ rotation and for proton pumping in the FoF1-ATPase. Both rotation and proton pumping were supported by the purine nucleotides, ATP, GTP, and ITP, but not by the pyrimidine nucleotides, CTP and UTP. Our results are consistent with the report by Perlin et al. (18Perlin D.S. Latchney L.R. Wise J.G. Senior A.E. Biochemistry. 1984; 23: 4998-5003Crossref PubMed Scopus (56) Google Scholar), where GTP and ITP were shown to drive proton pumping in the FoF1-ATPase, and support the idea that nucleotide hydrolysis is coupled to proton pumping through mechanical rotation of the γ subunit. Our rotation assay has revealed that many of the rotational characteristics are common to all three purine nucleotides. (i) The rotary torque is constant around 40 pN·nm. (ii) The rotation consists of discrete 120° steps. (iii) Each 120° step is driven by the hydrolysis of one nucleotide molecule. (iv) Backward steps as fast as the forward ones occur occasionally. The first three points imply that the mechanical work done in a 120° step is about 80 pN·nm for all three nucleotides, and thus the energy conversion efficiency can reach ∼100% with all three purine nucleotides. As to the last statement, we have shown in this study that the mechanical work of ∼80 pN·nm is done even when ΔG ATP or ΔG GTP is reduced to −80 pN·nm/molecule. We did not ascertain a high efficiency in ITP-driven rotation, but Sorgatoet al. (19Sorgato M.C. Galiazzo F. Valente M. Cavallini L. Ferguson S.J. Biochim. Biophys. Acta. 1982; 681: 319-322Crossref PubMed Scopus (4) Google Scholar) have reported that the energy from ITP hydrolysis is converted into a membrane potential by submitochondrial particles as efficiently as ATP hydrolysis. Thus, the efficiency of ITP-driven rotation is expected also to be close to 100%. The similarities among the three nucleotides suggest that the mechanical characteristics of the rotation such as the stepping, torque, and work per step are inherent in the (structure of the) F1motor. Purine nucleotides can trigger and let proceed the stepping mechanism in which the step angle, torque, and work are preset, whereas pyrimidine nucleotides cannot. Chemical kinetics, in contrast, are different among the three purine nucleotides. First, the hydrolysis curves in Fig. 3c are shifted toward higher concentrations in the order of ATP, GTP, and ITP, and the rotation curves follow the same trend. Apparently, this is due to the difference in the rate of nucleotide binding, ATP being the fastest and ITP the slowest. Analysis of step intervals at low nucleotide concentrations supported this view; the rate constant was 2.5 × 107m−1s−1 for ATP (10Yasuda R. Noji H. Kinosita Jr., K. Yoshida M. Cell. 1998; 93: 1117-1124Abstract Full Text Full Text PDF PubMed Scopus (713) Google Scholar), 2.9 × 106m−1 s−1 for GTP, and 1.1 × 106m−1 s−1 for ITP. According to the binding change mechanism for ATP synthase (1Boyer P.D. Biochim. Biophys. Acta. 2000; 1458: 252-262Crossref PubMed Scopus (98) Google Scholar), much of the energy available from γ rotation during synthesis is used to release a tightly bound ATP into the medium. Conversely, during hydrolysis, binding of ATP powers the reverse rotation of γ. If the above difference in the rate constant of nucleotide binding reflects a similar difference in the affinity for the nucleotide, then the energy provided by ITP binding will be the smallest and might have been insufficient to power the stepping mechanism. Because all three purine nucleotides supported rotation, it seems either that the affinity for ITP is high enough (a 20-fold difference in the binding constant is equivalent to a free energy difference of only 12 pN·nm/molecule), or that the rate of nucleotide release decreases in the order of ATP to ITP and makes the affinities more or less the same. In this regard, it is possible that UTP, of which the hydrolysis curve in Fig.3c is further shifted toward the right, cannot confer sufficient binding energy to the stepping mechanism. A second difference in the chemical kinetics is that the ATP hydrolysis was described by two sets of K m andV max, whereas one set sufficed for GTP and ITP. The two sets for the case of ATP are usually ascribed to two modes of catalysis: bisite and trisite, where one or two, or two or three, respectively, of the three catalytic sites are alternately filled with a nucleotide. The possibility, however, exists that the enzyme operates in the bisite mode at all concentrations at or above micromolar and that the apparent deviation from the simple Michaelis-Menten kinetics is due to the Mg-ADP inhibition. Single K m values for the hydrolysis of GTP and ITP, which are shown to be less prone to inhibition, support this view, although two sets ofK m and V max with similarV max/K m could also explain the simple kinetics. Resolution calls for the determination of the number of nucleotides bound in the catalytic sites of uninhibited enzyme, which is not an easy experiment. The pyrimidine nucleotides, CTP and UTP, were poor substrates for the TF1 and TFoF1. CTP was not hydrolyzed and, naturally, did not drive γ rotation or proton pump. UTP was hydrolyzed by TF1, yet UTP did not support rotation and pumping. Ca-ATP has also been reported to be an uncoupling substrate for FoF1, in that Ca-ATP was hydrolyzed without pumping protons (20Papageorgiou S. Melandri A.B. Solaini G. J. Bioenerg. Biomembr. 1998; 30: 533-541Crossref PubMed Scopus (36) Google Scholar). The kinetics of Ca-ATP hydrolysis, however, was similar to that of Mg-ATP hydrolysis. Thus, the nature of uncoupling seems different between the two cases, Ca-ATP and (Mg-)UTP. Interestingly, the reconstituted TFoF1 did not hydrolyze UTP, while TF1 (the α3β3γ subcomplex) did. This finding is consistent with the report by Yokoyama et al. (21Yokoyama K. Hisabori T. Yoshida M. J. Biol. Chem. 1989; 264: 21837-21841Abstract Full Text PDF PubMed Google Scholar) that the substrate specificity of this enzyme is higher when it contains a fuller complement of subunits. The UTP hydrolysis activity of the α3β3γ subcomplex may be ascribed to some flexibility in and around the catalytic site that would be suppressed in FoF1; such hydrolysis may proceed without a major structural change of the β subunit and thus without rotation of the γ subunit. We thank Y. Harada, T. Nishizaka, K. Adachi, T. Hisabori, and E. Muneyuki for technical assistance and helpful discussions and H. Umezawa for laboratory management." @default.
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• W2154767191 title "Purine but Not Pyrimidine Nucleotides Support Rotation of F1-ATPase" @default.
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