INTRODUCTION

ATP synthase (F₀F₁-ATPase), the enzyme that synthesizes and hydrolyzes the -phosphate bond of ATP, is crucial for the life of aerobic organisms. The enzyme resides in the inner mitochondrial membrane of animals, plants, yeast, and Neurospora; in the cytoplasmic membrane of bacteria and in the thylakoid membrane of chloroplasts in plants (see Refs. 1, 2, 3, 4, 5, 6 for reviews). In accordance with the chemiosmotic hypothesis (7) the electrical energy of respiration is first conserved as a protonmotive force (_H⁺), which via the F₀ moiety (8) of the synthase delivers the accumulated energy to the -subunits of the F₁ moiety (₃₃), presumably through conformational changes in the stalk. There is a wealth of experimental evidence supporting the view that the step with the largest demand for energy is the one involved in the release of ATP from the catalytic sites of the enzyme. According to basic postulates of the "binding change" mechanism (9, 10, 11), there is an equivalent participation of the three -subunits in the synthesis of ATP as they proceed through a cycle of "open," "loose," and "tight" states. Thus, at any one time, all three catalytic sites are in different conformations, but all pass sequentially through the same conformations (9, 10, 11). The alternating participation of catalytic sites was shown, for the first time by Adolfsen and Moundrianakis (12) in hydrolytic reactions catalyzed by bacterial F₁-ATPase.

Although the equivalent, alternating participation of catalytic sites within ATP synthases does not require positive cooperativity among them, many investigators believe that such cooperativity does in fact occur. The genesis of this long held view derives primarily from studies (13, 14) in which the F₁ moiety of the bovine heart enzyme was compared under unisite and multisite catalytic conditions, i.e. conditions in which the ATP/F₁ ratio was adjusted so that either one or all sites were operating. The resultant multisite/unisite rate enhancement ratio of 10⁶ was interpreted as reflecting strong positive cooperativity among catalytic sites (13). The bovine heart F₁ preparation used in these experiments contained bound nucleotide (2.8-3.5 mol/mol of F₁) and was prior incubated with ATP (13).

In studies reported here, and for the first time, the kinetics of ATP hydrolysis catalyzed by nucleotide-depleted F₁ have been determined by measuring the entire time course of individual reactions from the moment ATP (in the range from 1 nM to ~ 20 µM) enters in contact with the catalytic sites until it is totally hydrolyzed. Thus, catalytic sites are open when the experiment is commenced, and data are collected throughout the entire range from unisite through multisite reaction conditions. This study demonstrates that regardless of the ATP/F₁ molar ratio and enzyme source, bovine or rat, that all catalytic sites participating in ATP hydrolysis within F₁ can function in a kinetically equivalent manner.

EXPERIMENTAL PROCEDURES

Bovine heart F₁ was obtained from Drs. William Allison and J. M. Jault (University of California, San Diego). The enzyme was prepared by a modification (15) of the procedure of Knowles and Penefsky (16) and depleted of nucleotides as described by Garrett and Penefsky (17). It was stored prior to use in the refrigerator at 4 °C in 100 mM Tris-Cl, pH 8.0, 4 mM EDTA, and 50% glycerol (v/v). Rat liver F₁ was purified by a modification (18) of the procedure of Catterall and Pedersen (19). The purified enzyme, in 250 mM KP_i and 5.0 mM EDTA, was divided into 100-µl aliquots and lyophilized to dryness and stored at 20 °C until use. Immediately before use the enzyme was redissolved in 100 µl of water and precipitated twice with ammonium sulfate. The bovine heart and rat liver F₁ preparations contained, respectively, <0.04 (11 determinations) and 0.9 (7 determinations) tightly bound nucleotide/mol enzyme when determined after denaturation by the highly sensitive chemiluminescent assay described below. As it is known that such preparations can bind a total of 5-6 mol of nucleotide/mol of F₁ (20, 21), these preparations are appropriately defined as "nucleotide-depleted." In fact, the bovine heart F₁ preparation employed here is essentially nucleotide free.

ATP was a product of Sigma and the 1243-200 ATP monitoring reagent, a mixture of luciferin and luciferase was a product of BioOrbit. The luminometer used in these studies to monitor ATP disappearance was a product of LKB (Wallac model 125), and the attached recorder was a product of Soltec (model 330). [-³²P]ATP was purchased from DuPont NEN and its radioactivity assessed in a Beckman LS600IC liquid scintillation counter using Budget solve complete counting mixture (Research Products International Corp.). Activated charcoal (number C4386), prewashed with HCl, was a product of Sigma, and the filtration device (Centricon 100, molecular weight cutoff = 100,000) used to separate F₁ and charcoal-bound [-³²P]ATP from ³²P_i was obtained from Amicon.

The standard reaction medium, in 1.0 ml final volume at 24 °C, consisted of 200 mM sucrose, 50 mM KCl, 10 mM NaP_i, pH 7.05, 2 mM MgS0₄, and 50 µl of a 5-ml solution in distilled water of the ATP monitoring reagent. ATP in amounts indicated in the legends to the tables and figures was then added to the stirred reaction medium to elicit the chemiluminescent response, followed by the addition of F₁ to initiate the ATPase reaction. The reverse (adding ATP to suspensions of F₁) was also possible without affecting the steady-state kinetics of the reaction. Changes in ATP concentration were monitored with an LKB Wallac model 125 luminometer. The electrical signal elicited by light emission was suitably amplified up to 10,000 times by changes in the current (from 10⁶ to 10⁸ A) and the voltage (from 10 V to 1 mV), and recorded using a Soltec model 330 multichannel recorder, usually run at a chart speed of 120 cm/min. The contents of the cell were stirred with a magnetic bar rotating at speeds of near 1000 rpm by means of an electrical device placed on the side of the reaction chamber. The concentration of ATP, in different standard solutions, was spectrophotometrically determined from the absorbance at 259 nm using a millimolar extinction coefficient of 15.4.

As the observed initial rates of decrease of the chemiluminescent signal were always first order with respect to the concentration of ATP (d[ATP]/dt = k[ATP]), the rate constant "k," was calculated from the integrated form of the equation (ln[ATP]_t = ln[ATP]₀  kt) by plotting 1n[ATP]_t versus t at 400-ms intervals. Only steady-state segments (>70% of the reaction) were considered in these calculations (Fig. 2).

The entire reaction was carried out in a Centricon-100 filtration device. The standard reaction medium, again in 1-ml final volume at 24 °C, was identical to that described above except that the ATP monitoring reagent was omitted and [-³²P]ATP (2.1 × 10⁶ cpm) rather than "cold" ATP was included in the assay. The reaction was initiated with F₁, quenched at 2 s by addition of 1 ml of 100 mg/ml activated charcoal, and immediately centrifuged at 3000 rpm (1086 × g) in a SS-34 rotor in a Sorvall RC-2B centrifuge at 4 °C. The filtrate containing ³²P_i released in the ATPase reaction was then assessed for radioactivity in 10 ml of Budget Solve as indicated above.

Protein was determined by the method of Lowry et al. (22) after first precipitating with 5% trichloroacetic acid.

RESULTS AND DISCUSSION

In kinetic studies described below ATP hydrolysis was monitored by following the disappearance of the chemiluminescent signal induced by adding ATP to an assay medium containing luciferin and luciferase. Prior to commencing these experiments, it was important to define the response time of the system for detecting ATP and to establish whether concentrations of ATP could be accurately detected at ratios of ATP/F₁ in the assay ranging from less than 1 to much greater than 1 (i.e. from unisite to multisite conditions). Confirming previous studies of DeLuca et al. (23, 24), the results depicted in Fig. 1A (traces a to c) show that, regardless of the amount of ATP (1-20,000 pmol), the response time of the luciferin-luciferase system is close to 300 ms, much faster than the time course of the ATPase reaction at concentrations of ATP and F₁ used in this study. The accuracy of the technique depicted in Fig. 1B, the data of which were derived from over 70 different experiments, shows that the correlation coefficient between light emission and ATP concentration is better than 0.999 at ATP concentrations of 15-17 µM ATP. As very low F₁ concentrations are used in the studies described below, the technique allows for accurate ATP measurements to be made with assays in which the ATP/F₁ ratio ranges from ~0.1 to ~1900. Above 15-17 µM ATP, the extent of the chemiluminescent signal rapidly decreases as the concentration of ATP is increased (Fig. 1), due most likely to the accumulation of dehydroluciferin (23). For this reason, and for the reason indicated below, the hydrolytic reaction was initiated by injecting F₁ into the reaction cell already containing ATP.

Three conditions were adhered to in the performance of all experiments. First, nucleotide-depleted preparations of F₁ (bovine heart or rat liver) were used to avoid any possible allosteric effects resulting from nucleotides bound to noncatalytic sites, i.e. sites located predominantly on -subunits (25). Second, the reaction was initiated by adding F₁ to the reaction mixture already containing ATP to allow the nucleotide free catalytic sites to bind and hydrolyze ATP immediately upon contacting ATP. Third, and in contrast to the earlier studies (13, 14), prior incubation of F₁ with ATP ("aging") was avoided both to allow for detection of the actual initial rates of ATP hydrolysis and to avoid product (ADP) inhibition that might give rise to "apparent" cooperative kinetics upon addition of excess ATP.

Fig. 2 shows that under the above conditions the initial rates of ATP hydrolysis depend on ATP concentration to the first power as precisely defined by the first order rate equation d[ATP]/dt = k[ATP]. This is true regardless of the source of enzyme (bovine heart or rat liver) or whether the reaction takes place under "unisite" (ATP/F₁ = 0.02, Fig. 2A) or "multisite" (ATP/F₁ = 48.5, Fig. 2B) catalytic conditions. Plots (insets in Fig. 2) of the integrated form of the first order rate equation (ln[ATP]_t = ln[ATP]₀  kt) were used to calculate the first order rate constant, k, which in turn was used to calculate the turnover number. For example, for the hydrolysis of 1 pmol of ATP by bovine heart F₁ (45.9 pmol) the first order rate constant, k, obtained from the plot ln[ATP] versus t (Fig. 2A, inset) was 0.475 s¹, and the calculated turnover number (1 pmol of ATP × 0.475 s¹/45.9 pmol of F₁) was 0.010 s¹.

It has been reported that the F₁ moiety of bovine heart F₁ catalyzes the hydrolysis of ATP by a mechanism in which the slow rate of hydrolysis at a single catalytic site is enhanced by ~10⁶-fold when the ATP concentration is increased by more than 3 × 10³-fold, a process interpreted to result in strong cooperative interactions between catalytic sites (13). The important data leading to this conclusion is actually derived from two separate experiments. The first was conducted under (see Fig. 5 in Ref. 14) in which a turnover number of was determined by monitoring the dissociation of ADP remaining bound to F₁ after prior incubation (aging) of ATP and excess F₁ (ATP/F₁ = 0.2) The second experiment was conducted under multisite conditions (Fig. 1 in Ref. 13) where a turnover number of was determined by monitoring release of ³²P_i from [-³²P]ATP after adding ATP (2.5 ml, 10 mM) in excess to a unisite mixture of F₁ (3 µM in 2.5 ml) and [-³²P]ATP (0.3 µM in 2.5 ml) that had been prior incubated for 2 s. (The multisite turnover number is somewhat higher ( ) when measured by the authors under steady-state conditions in the presence of an ATP regenerating system.) The was calculated from the simple multisite/unisite ratio of 300 (or 600 s¹)/3.6 × 10⁴ s¹.

In experiments reported here (Tables I and II, Fig. 3), and in contrast to those described above, nucleotide-depleted F₁ preparations (bovine and rat) were employed; a single method (chemiluminescent assay) was used to measure both unisite and multisite turnover rates; the entire time course of the reaction was monitored at each ATP concentration as indicated in Fig. 2; and ATP concentration was varied over a wide range. Table I and Table II summarize kinetic data obtained for typical experiments with bovine and rat liver F₁, respectively. The first order rate constant for ATP hydrolysis remains relatively constant under both unisite and multisite conditions declining only slightly at high concentrations of ATP. Turnover numbers under unisite conditions range from at the lowest ATP/F₁ assay ratios, i.e. from 0.11 to 0.55. Fig. 3, A and B, summarize and versus (Lineweaver-Burk) plots of the data presented in Table I for bovine heart F₁. Significantly in neither plot, which spans the range from unisite condition, with an ATP/F₁ assay ratio as small as 0.11, to multisite conditions, with an ATP/F₁ ratio as high as 1900, is there any indication of sigmoidicity characteristic of cooperative behavior. Rather, it is clear that bovine heart F₁ assayed under these conditions exhibits strictly Michaelis-Menten kinetic behavior consistent with the view that all participating catalytic sites are kinetically equivalent. The extrapolated single K_m is 57 µM, and the extrapolated V_max is 103 µmol of ATP hydrolyzed per min/mg of F₁. The turnover number of 635 s¹ is very close to the value of 600 s¹ obtained under multisite conditions in the earlier study (Ref. 13, see discussion above). Similar results were obtained with rat liver F₁ (Table II, Fig. 3B), which exhibits a V_max nearly identical to bovine heart F₁ and a slightly higher K_m of 79 µM.

Table I. Dependence of the rates of ATP hydrolysis by bovine heart F1 on ATP concentration Reactions were initiated by injecting 9.16 pmol of nucleotide-depleted bovine heart F1 into a 1 ml chemiluminescent reaction system containing the indicated amount of ATP. Assays were carried out exactly as described under "Experimental Procedures." The first order rate constant (k) of the reaction was calculated from the slope of the line that results from plotting the integrated form of the first order rate equation: dATP/dt = k[ATP] at 0.4-s intervals. A molecular mass for F1 of 371,000 was used in calculating turnover numbers. Values represent averages of duplicate determinations. ATP added First order rate constant Turnover number Specific activity ATP hydrolyzed pmol     s1 nmol/min/mg protein 1 0.103 0.011 1.81 2 0.103 0.022 3.62 5 0.103 0.056 9.07 10 0.102 0.111 17.92 20 0.103 0.225 37.01 50 0.102 0.560 90.03 100 0.102 1.111 179.69 200 0.102 2.223 359.52 500 0.102 5.559 899.07 1000 0.101 11.009 1780.42 1710 0.100 18.639 3014.34 4275 0.096 44.803 7237.75 8550 0.090 83.875 13,505.76 17,100 0.080 149.11 24,109.96

Table II. Dependence of the rates of ATP hydrolysis by rat liver F1 on ATP concentration Reactions were initiated by injecting 29.6 pmol of nucleotide-depleted F1 into a 1-ml chemiluminescent reaction system containing the indicated amount of ATP. Assays were carried out exactly as described under "Experimental Procedures." The first order rate constant (k) of the reaction was calculated from the slope of the lines that result from plotting the integrated form of the first order rate equation dATP/dt = k[ATP] at 0.4-s intervals. Values represent averages of duplicate determinations. ATP added First order rate constant Turnover number Specific activity ATP hydrolyzed pmol     s1 nmol/min/mg protein 2 0.237 0.016 2.532 5 0.237 0.040 6.319 7.5 0.237 0.060 9.487 10 0.237 0.080 12.628 15 0.237 0.120 18.974 20 0.237 0.160 25.277 25 0.237 0.200 31.610 50 0.237 0.400 63.193 100 0.237 0.798 126.01 250 0.237 1.997 315.30 500 0.236 3.976 627.67 1000 0.234 7.885 1244.72 2500 0.230 19.377 3058.99 5000 0.223 37.575 5931.79 10000 0.211 71.107 11,225.41

Comparison of the Chemiluminescent Assay for Monitoring ATP Hydrolysis with an Assay That Monitors Release of ³²P_i from [-³²P]ATP

Although the turnover number of 635 s¹ for multisite catalytic conditions reported here for the bovine heart F₁ is nearly identical to that reported in the earlier study (13), the unisite turnover numbers at the lowest ATP/F₁ ratios are much higher, in the range of 0.01-0.06 s¹ (Tables I and II), rather than near 10⁴ s¹ (13, 14). Consequently, data obtained here result in multisite/unisite ratios near 10⁴ rather than near 10⁶ (13) as reported earlier. For this reason, it might be argued that the chemiluminescent assay, which monitors ATP disappearance, may not accurately report ATP hydrolytic rates at low ATP/F₁ assay ratios (unisite conditions). To address this question, we compared the chemiluminescent assay for monitoring ATP hydrolysis under unisite conditions with that of an assay that monitors ³²P_i release from [-³²P]ATP. Two experimental conditions were chosen, one in which the ATP/F₁ assay ratio was only 0.11 (lowest data point in Table I), and one in which the ratio was 1.1. The assay medium was identical to that used for monitoring ATP disappearance except that luciferin and luciferase were omitted. Significantly, the specific activities of 2.9 ± 0.21 and 14.5 ± 2.9 nmol of P_i released per min/mg of protein obtained using the ³²P_i release assay (Table III) compare favorably with those of 1.8 and 18 nmol of ATP disappeared per min/mg of protein obtained using the chemiluminescent assay (Table I), for ATP/F₁ ratios, respectively, of 0.11 and 1.1. Therefore, it seems clear that the chemiluminescent assay is a reliable indicator of the ATP hydrolytic rate under unisite as well as multisite conditions.

Table III. ATP hydrolysis catalyzed by bovine heart F1 under unisite conditions using a radioactive assay for monitoring the release of 32Pi from [-32P]ATP Reaction conditions were identical to those used in the chemiluminescent assay (see "Experimental Procedures") except that the luciferin-luciferase reagent was omitted and [-32P]ATP rather than "cold" ATP was present. Upon addition of 9.16 pmol of nucleotide-depleted bovine heart F1 to the assay medium contained within a Centricon-100 filtration device, the reaction was allowed to proceed for 2 s followed by immediate quenching with activated charcoal. Separation of 32Pi from charcoal bound [32P]ATP was performed by centrifugation (see "Experimental Procedures"). Specific activity values are reported as mean ± the standard deviation. ATP added ATP/F1 in assay Number of experiments Specific activity pmol nmols Pi/min/mg protein 1 0.11 4 2.9  ± 0.21 10 1.1 10 14.5  ± 2.9

In studies described here, the bovine heart F₁ preparation was essentially nucleotide-free, and the rat liver F₁ preparation contained less than 1 mol/mol of F₁. Both enzymes exhibited strictly Michaelis-Menten kinetic behavior with maximal turnover numbers of 635 s¹ and multisite/unisite rate enhancement ratios near 10⁴, consistent with the view that F₁-ATPases can catalyze ATP hydrolysis by a mechanism in which all participating catalytic sites are kinetically equivalent. In earlier studies (13, 14), the bovine heart F₁ preparations used contained 2.8-3.5 mol of bound nucleotide/mol of F₁. These preparations exhibited maximal turnover numbers of 600 s¹ and multisite/unisite rate enhancement ratios of 10⁶, consistent with the view that F₁-ATPases can catalyze ATP hydrolysis by a mechanism in which positive cooperativity occurs between catalytic sites. Perhaps the simplest interpretations of the two studies is that, depending on nucleotide content and its subunit distribution, F₁ ATPases can exist in different conformational states, one in which all participating catalytic sites are kinetically equivalent and one in which they are kinetically nonequivalent. In support of this view are two different x-ray structures of F₁ (25, 26), one in which the catalytic -subunits appear more structurally equivalent (25) than in the other (26).

Alternative interpretations are possible and require further investigation. One possibility is that F₁-ATPases normally function during ATP hydrolysis as simple Michaelis-Menten enzymes and that deviations from this behavior (i.e. sigmoid kinetic behavior), rather than reflecting positive catalytic cooperativity, simply reflect inhibitory ADP bound at a catalytic site that is displaced as ATP concentration is raised. As shown in Fig. 4, ADP is a potent inhibitor of bovine heart F₁ with over 90% inhibition being observed when the ADP concentration equals the ATP concentration at values 5 µM. Significantly, F₁ preparations are frequently stored or isolated in ATP, the hydrolysis of which results in F₁·ADP complex formation.

Specifically, as it applies to the earlier studies (13, 14) leading to the view that bovine heart F₁ exhibits positive catalytic cooperativity, it is interesting to note that the unisite turnover number of 10⁴ s¹ was based neither on the release of P_i nor ADP following ATP hydrolysis per se, but rather on the dissociation of ADP remaining bound to F₁ following hydrolysis (Fig. 5 in Ref. 14). Thus, the possibility exists that the unisite turnover number of 10⁴ s¹ obtained from this earlier analysis is not an accurate reflection of the actual value and corresponds to release of ADP from a noncatalytic site. It is important to note that the same investigators in a separate study (Fig. 1 in Ref. 13) find that incubation of [-³²P]ATP (0.3 µM in 2.5 ml) under unisite conditions with F₁ (3 µM in 2.5 ml) for 2 s prior to adding excess ATP results in 20% hydrolysis of the total ATP present. Had these investigators calculated the unisite turnover number under these conditions, they would have obtained a value of 0.01 s¹ (i.e. 0.15 µM ATP × 0.20/1.5 µM F₁ × 2 s). This value is almost identical to the unisite turnover number of 0.011 s¹ reported here as the first entry in Table I and consistent with Michaelis-Menten rather than cooperative kinetic behavior.

Finally, consistent with the studies reported here, recent work on nucleotide-depleted Escherichia coli F₁ (27) show that in the ATP concentration range of 1 µM to 1 mM a single K_m value of 38 µM is sufficient for an adequate description of the ATP hydrolytic behavior of the enzyme. However, these authors believe that all catalytic sites (presumably three) must be occupied to achieve significant rates of ATP hydrolysis. Clearly, this is not the case with the nucleotide-depleted F₁ preparations from the animal systems examined here, as significant rates of ATP hydrolysis are readily detected at ATP/F₁ assay ratios as low as 0.02 (Fig. 2A). Moreover, at 17 µM ATP, well below the K_m of 57 µM, the catalytic turnover number is already 149 s¹ (~24 µmol of ATP hydrolyzed per min/mg of F₁), as shown in Table I. Thus, nucleotide-depleted animal F₁ preparations, in accordance with simple Michaelis-Menten kinetic behavior, show significant rates of ATP hydrolysis at all concentrations of ATP tested.