Effect of the Lipid Environment on Protein Motion and Enzymatic Activity of the Sarcoplasmic Reticulum Calcium ATPase*

Transcription

1 Effect of the Lipid Environment on Protein Motion and Enzymatic Activity of the Sarcoplasmic Reticulum Calcium ATPase* (Received for publication, April 4, 1978) Cecilia Hidalgo, David D. Thomas,+ and Noriaki Ikemoto From the Department of Muscle Research, Boston Biomedical Research Institute, Boston, Massachusetts 02114, and the Department of Neurology, Harvard Medical School, Boston, Massachusetts In order to investigate the roles of the physical states of phospholipid and protein in the enzymatic behavior of the Ca +-ATPase from sarcoplasmic reticulum, we have modified the lipid phase of the enzyme, observed the effects on the enzymatic activity at low temperatures, and correlated these effects with spectroscopic measurements of the rotational motions of both the lipid and protein components. Replacement of the native lipids with dipalmitoyl phosphatidylcholine inhibits ATPase activity and decreases both lipid fluidity, as monitored by EPR spectroscopy on a stearic acid spin label, and protein rotational mobility, as monitored by saturation transfer EPR spectroscopy on the covalently spin-labeled enzyme. Solubilization of the lipid-replaced enzyme with Triton X-100 reverses all three of these effects. Ten millimolar CaClz added either to the enzyme associated with the endogenous lipids or to the T&on X-100 solubilized enzyme inhibits both ATPase activity and protein rotational mobility but has no detectable effect on the lipid mobility. These results are consistent with the proposal that both lipid fluidity and protein rotational mobility are essential for enzymatic activity. The evidence accumulated over the last several years indicates that phospholipids are essential for both the ATPase reaction and the coupled calcium transport in sarcoplasmic reticulum (SR). Partial removal of phospholipids results in complete inhibition of ATP hydrolysis, with little or no effect on the formation of a phosphorylated enzyme intermediate (1, 2), indicating a stringent phospholipid requirement in the reaction step(s) in which the decomposition of phosphoenzyme takes place, rather than in the phosphoenzyme forma- * This research was supported by grants from the American Heart Association (74913), the National Institutes of Health (AM 16922, HL-05811, HL-05949, and RR-05711), the National Science Foundation, and the Muscular Dystrophy Associations of America, Inc. 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 USC. Section 1734 solely to indicate this fact. $ During the course of this work, D. D. T. was a Postdoctoral Fellow of the Muscular Dystrophy Associations of America. Present address, Department of Structural Biology, Stanford University School of Medicine, Stanford, Calif I The abbreviations used are: SR, sarcoplasmic reticulum; DPL, dipalmitoyl lecithin; SR-ATPase, purified Ca +-ATPase containing the endogenous SR lipids; DPL-ATPase*, partially purified Ca *- ATPase containing DPL; DPL-ATPase, purified Ca*+-ATPase containing DPL. MSL, maleimide spin label, N-( l-oxyl-2,2,6,6-tetramethyl-4-piperidinyl)maleimide; MalNEt, N-ethylmaleimide; EGTA, ethylene glycol bis(p-aminoethyl ether) N,N -tetraacetic acid; DNP, 2,4-dinitrophenyl. tion step. It has been recently reported that the Ca +-ATPase is active for several days in true solution in the presence of some nonionic detergents (3). The soluble enzyme retains about 30 mol of tightly bound phospholipid per mol of polypeptide chain, indicating the existence of strong attractive forces between protein and lipid. Removal with ionic detergents of most of the membrane phospholipid results in substantial inhibition of phosphoenzyme formation, which can be reversed by the subsequent addition of either phosphatidylcholine (4) or the nonionic detergent dodecyl oxyethyleneglyco1 monoether (5). This suggests that simpler amphiphiles can replace phospholipids in restoring ATPase activity. However, complete delipidation by deoxycholate results in irreversible inactivation (6). New possibilities for the study of the involvement of phospholipids in the ATPase reaction have been opened by the development of procedures which allow the replacement of the endogenous phospholipids of SR by chemically well defined phospholipid species (7). It has been shown that replacement with a saturated lecithin, dipalmitoyl phosphatidylcholine (DPL), completely inhibits the Ca2+-ATPase activity at low temperatures (7-9). Furthermore, studies on the formation and decomposition of the phosphorylated intermediate have shown that replacement with DPL strongly inhibits the rate of ATP hydrolysis by inhibiting the decomposition of the phosphorylated intermediate, with little or no effect on the phosphorylation reaction and its reversal (8,9). The inhibition takes place at temperatures at which the mobility of the lipid hydrocarbon chains is strongly restricted (8), indicating that the physical state of the phospholipids associated with the Ca2 -ATPase enzyme is an important factor in determining the rate of ATP hydrolysis in SR. Parallel studies in SR of the effect of temperature on enzymatic activity, lipid fluidity, and mobility of protein-bound spin labels have indicated a correlation among these variables. At the transition temperature observed in Arrhenius plots of the Ca2+-ATPase activity, breaks have been obtained in plots of the effect of temperature on spectral parameters obtained either from protein-bound spin labels (10) or from spin labels incorporated into the lipid phase (8, 10). All of this clearly indicates that the action of the Ca2+-ATPase enzyme, and presumably of other membrane-bound enzymes, involves the dynamic interactions of proteins and lipids. The present work is an attempt to further characterize the mechanism by which the physical state of the phospholipids affects the enzymatic activity of the Ca2+-ATPase from SR. The experimental approach followed in this work was to modify the lipid phase of the Ca +-ATPase enzyme, observe the effects on the enzymatic activity, and correlate these effects with direct EPR measurements of the molecular motions of both the lipid and the protein components. Two EPR techniques were used: the conventional EPR technique, in- 6879

2 Role of Protein and Lipid Motion in Ca -ATPase Activity valving measurement of first derivative spectra; and the saturation transfer technique, which allows determinations of considerably slower rotational motions (ll), such as the motion of a membrane-bound protein as a whole or of large segments within the protein. The results indicate that, at low temperatures, replacement of the endogenous SR lipids with DPL, in addition to the strong inhibition of ATPase activity and the decrease in lipid mobility previously described (8), produces a considerable decrease in protein motion. Addition of Triton X-100 to the DPL-replaced enzyme preparations, resulting in enzyme solubilization, restores enzymatic activity and produces a considerable increase in the mobility of both the DPL molecules and the protein. These findings indicate that the constraints imposed by the DPL-containing membrane structure on its components are effectively removed by the detergent, allowing the enzyme to hydrolyze ATP at the normal rate. The strong inhibition of the Ca2 -ATPase reaction by [Ca ] in the millimolar range, resulting from inhibition of the decomposition of the phosphorylated intermediate in both the membrane-bound (containing the endogenous SR lipids) and the soluble forma of the Gas+-ATPaae enzyme, ia accompanied by a reduction in the mobility of the protein but not of the lipids. The selective reduction of protein motion induced by [Ca ] in the millimolar range is probably not due to formation of protein protein or protein.lipid complexes, as determined by gel filtration chromatography of the soluble enzyme with or without 10 mm Ca +. These results are consistent with a model of the Caz+-ATPase enzyme in which a certain degree of protein and lipid motion is required for enzymatic activity. EXPERIMENTAL Membrane PROCEDURES Preparations Preparation of fragmented SR from rabbit white skeletal muscle, purification of the Ca +-ATPase, and replacement of the endogenous membrane lipids by DPL were carried out as described previously (8, 12). A DPL-replaced ATPase preparation obtained from SR, previously designated DPL enzyme (8), contains a major loo,ooo-dalton protein band, with a few minor components of lower molecular weights as revealed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis. In this paper, it will be referred to as DPL-ATPase*. The purified ATPase enzyme, containing a single KQOOO-dalton protein component plus the endogenous SR lipids, and the DPLreplaced preparation, obtained from it by using the same replacement procedure described for SR (8), will be referred to as SR-ATPase and DPL-ATPase, respectively. Both DPL-replaced ATPase preparations contain about 30 mol of DPL/10 g of protein. This was determined by adding a small amount of [14C]DPL to the initial deoxycholate- DPL solution used in the replacement procedure, measuring the specific radioactivity of the DPL solution, and determining the content of [14C]DPL of the resulting membrane fractions. Deoxycholate, Triton X-190, and L-OL-DPL were obtained from Sigma Chemical Co. and [ C]DPL from Scientific Products. Deoxycholate was recqstallized prior to use; both DPL and Triton X-106 were used without further purification. Enzymatic Assays Determination of ATPase activity, phosphoenzyme formation, and phosphate liberation was done as described previously (8), using [v- P]ATP obtained from New England Nuclear. To determine the effect of Triton X-100 on enzymatic activity, the protein was incubated with Triton for 2 min before the start of the reaction at the concentration stated (see figure and table legends). To measure the formation of phosphoenzyme as a function of [Ca* ], Ca2 -EGTA buffers were used; the Ca + concentrations were calculated from published Ca*+ and H+ binding constants with the use of a computer program (13). To assure complete equilibration, the protein was preincubated in the buffered Ca2+ solutions for 20 min at 0 C before starting the reaction by the addition of [T-~ P]ATP. Protein concentrations were determined by the method of Lowry et al. (14). Gel Chromatography Columns (1.5 x 90 cm) of Bio-Gel A-5m (Bio-Rad) were used. Blue dextran and CaCIZ were used as void and total volume markers, respectively. Ten milligrams of protein in a volume of 1 ml was placed on the column, which had been previously equilibrated with the solution used for elution. Fractions of 1.25 ml were collected at a flow rate of about 3 ml/h. EPR Measurements Spin Labeling-A stearic acid N-oxyl-4,4 -dimethyloxazolidine compound with the formula: CH:&CHZ),T-C-(CH&-COOH /\ 9,N-0 Y was used as a spin probe for the lipid phase. It was dissolved in benzene at a concentration of 10 nuu and stored at -20 C. The spin label was incorporated into the membrane as described previously (8), with a final ratio of one spin label per 100 molecules of membrane phospholipid. A maleimide spin label derivative, N-(l-oxyl-2,2,6,6- tetramethyl-4-piperidinyl)maleimide, was used to label the enzyme polypeptide. A fresh solution of spin label in ethanol was diluted lofold with buffer and added to SR-ATPase or to DPL-ATPase (final ethanol concentration less than l%), at a ratio of 1.25 mol of label/l@ g of protein, and incubated overnight at 0 C. Labeling was carried out in 0.3 M sucrose, 20 mm Tris/maleate, ph 7.0. To stop the reaction, the incubation solution was diluted 30-fold with an ice cold solution of 0.3 M sucrose, 20 mm Tris/maleate, ph 7.0, and the labeled enzyme was sedimented by centrifugation at 100,000 x g for 1 h. The pellet was resuspended in 0.3 M sucrose, 20 mru Tris/maleate, ph 7.0, at a protein concentration of 10 to 20 mg/ml. The resulting preparations were denoted MSL-SR-ATPase and MSL-DPL-ATPase. As determined by double integration of EPR spectra using a Nicolet 1074 computer, the final product contained 1 to 1.2 mol of label bound/lo g of protein. We have recently shown (15) that a minor glycoprotein component of SR contains sulfbydryl groups that react readily with the maleimide spin label. As stated earlier, DPL-ATPase* contains other minor components besides the ATPase polypeptide. For this reason, to avoid contributions to the spectra arising from components other than the ATPase polypeptide, only SR-ATPase and DPL-ATPase were used to spin label the enzyme polypeptide. However, to provide continuity with our previous work (8), we used DPL-ATPase for enzymatic and for lipid spin label experiments; essentially the same results as with DPL-ATPase were obtained. Spectroscopy-EPR experiments were performed with a Varian E- 109E spectrometer in the absorption mode. Samples were contained in a small Varian quartz flat cell ( Variable Temperature Aqueous Cell ), and the temperature was controlled to within 0.5 C with a V4540 variable temperature controller. Conventional spectra, designated V, were recorded with both field modulation and phase-sensitive detection at 100 khz. The peak-to-peak modulation amplitude was 2 G and the microwave power setting was 10 milliwatts. Saturation transfer spectra, designated V 2, were obtained essentially as described previously (11). The magnetic field was modulated at 50 khz and the phase-sensitive detection was at 100 khz. The modulation amplitude was 6.3 G. The reference phase was set 90 out-ofphase by minimizing the signal in the absence of saturation (power setting 0.5 milliwatt). The incident microwave power was 34 milliwatts. Using peroxylamine disulfonate, as described previously (ll), we determined that this power setting corresponded to an effective microwave field strength of HI = 0.25 G inside the sample cell. Unless otherwise noted, all spectra were obtained from preparations in 0.3 M sucrose, 20 mm Tris/maleate, ph 7.0, at 4, containing 10 to 20 mg of protein/ml. Since there is no preferential orientation with respect to the magnetic field for the membranes of freely suspended vesicles, the geometric details of anisotropic molecular motions are not precisely * In other studies, we have improved the selectivity of spin labeling the ATPase in the intact SR membrane by an alternative method, blocking the fast reacting sulthydryl groups with MalNEt before spin labeling (15).

3 Role of Protein and Lipid Motion in Ca2 -ATPase Activity determined. In our results below, we characterize our spectra with the parameters 2T k and L /L (see Fig. 1). Effective correlation times can be estimated from these parameters with the use of the curve in Fig. 1, assuming that the rotational motions are isotropic. Since the motions are almost certainly not isotropic, these measurements provide good estimates of the approximate time ranges of the motions and are useful for comparative purposes, but should not be interpreted as accurate correlation times describing complex molecular motions. Note in Fig. 1 that the parameter 2T ll, from conventional spectra, is sensitive only to rotational correlation times (~2) less than IO- s, while L /L, from saturatidn transfer spectra, is sensitive to Q values as long as 1 ms. Therefore, we can use 2T ll to characterize the relatively rapid motions of fluid lipid hydrocarbon chains, small segments of proteins, and proteins in solution. 2T ll, and hence also 2T ll, are also sensitive to changes in the polarity of the environment of the spin label. In the absence of changes in 2T ll, when 2T ll approaches 2 2 1, we can use L /L to characterize the slower motions of lipid hydrocarbon chains in the gel phase, whole proteins, or large protein segments, assembled into restrictive structures such as membranes. The utility of the saturation transfer method has been demonstrated in previous studies on assemblies of muscle proteins (19). RESULTS Enzymatic Activity The addition of Triton X-100 to DPL-ATPase* results in progressive stimulation of the rate of Pi liberation (Fig. 2), from less than 0.5 nmol mg- min- in the absence of detergent to about 7 nmol mg- min- at a Triton X-loo/protein weight ratio of 1.5. Since this stimulation of the Pi liberation rate is accompanied by a decrease in the steady state phosphoenzyme concentration, there must be a considerable increase in the rate constant k,,, defined as the ratio of the Pi liberation rate over the steady state phosphoenzyme level (Fig. 2). The turbidity of the DPL-ATPase* suspension decreases as Triton X-100 is added. Complete solubilization was not obtained owing to the presence of minor protein contaminants in DPL- ATPase* which are insoluble in Triton. However, the loo,ooo r2 bxnds) FIG. 1. Plots used to determine effective correlation times (~2) from nitroxide spin label EPR spectra. Left (dashed line), (2ql - 2T ll) uersus 72 where 2Tll is the separation in gauss of the outer extrema of a conventional spectrum corresponding to no motion (p2 > 10 s), and 2T ll is this separation when the spin label undergoes isotropic Brownian rotational motion with a correlation time TV, as calculated in computer simulations (16). Right (solid line), L /L uersus TV, where L and L are two peaks in the saturation transfer spectrum of a spin label undergoing isotropic Brownian rotational motion with a correlation time T:! (see inset). Closed circles are from computer simulations (11, 17) and open circles are from experiments on spin-labeled Hb in aqueous glycerol solutions. At Q = m, the open square is from ammonium sulfate-precipitated Hb, the closed square is from crystallized hemoglobin, and the triangle is from a pellet of spin-labeled F-actin (18) I mg of Trdon X-100 / mg cd protmn FIG. 2. Effect of Triton X-100 at 0 C on DPL-ATPase* reflected in the formation of phosphoenzyme, the rate of P, liberation, and the rate of phosphoenzyme decomposition. The ATPase reaction mixture contained 0.1 M KCl, 5 mm MgCl2, 10 pm CaCh, 20 mm Tris/maleate, ph 7.0, and 10 pm [v- PI ATP. The reaction was started by addition of the Triton X-lOO/protein mixture, at the Triton/protein weight ratios indicated on the abscissa. Final concentration of protein, 0.2 mg/ml. The protein was incubated with Triton X-100 for 2 min at 0 C before starting the reaction; addition to the reaction solution resulted in a IO-fold dilution of the Triton/protein mixtures. The reaction was stopped by addition of 6.7% trichloroacetic acid. The amount of phosphoenzyme formed, the rates of P, liberation, and the reaction rate of phosphoenzyme decomposition were determined as described under Experimental Procedures. The results of two different experiments are shown separately for phosphoenzyme formation and for rate of P, liberation. The values for the rate constants are the averages between the two different preparations. 0, W, phosphoenzyme; 0, Cl, P, liberation rates; A, rate constant of phosphoenzyme decomposition. dalton ATPase peptide was solubilized, as shown by phosphoenzyme values >l nmol/mg of protein in the soluble fraction. In contrast, the phosphoenzyme levels of the Tritoninsoluble fraction were below 0.1 nmol/mg. DPL-ATPase, which contains only the ATPase polypeptide and DPL, is completely solubilized by Triton. The effect of Triton X-100 on Pi liberation rates is shown in Table I for DPL-ATPase*, DPL-ATPase, and SR-ATPase. Several features are to be noted. (a) In the absence of T&on X-100, the rates of Pi liberation in DPL-ATPase* and DPL- ATPase at 0 C are more than 10 times lower than in SR- ATPase, as described previously fpr DPL-ATPase* (8); (6) the addition of Triton X-100 to both DPL-ATPase* and DPL- ATPase produces a substantial increase, from lo- to 30-fold, in the rates of Pi liberation at a Triton/protein weight ratio of 1.5 (CL Fig. 2); (c) addition of the same amount of Triton X- 100 to SR-ATPase does not affect the rate of P, liberation, despite the fact that it causes complete solubilization of the enzyme; and (d) the addition of 10 mu CaClz results in 92% inhibition of the Pi liberation rate of SR-ATPase, regardless of the presence of Triton X-100. In both DPL-replaced preparations, the addition of 10 mu CaCl2 in the absence of Triton X-100 has little inhibitory effect on the already low DPLinhibited rates of Pi liberation. However, in the presence of Triton X-100 the enhanced ATPase rate of DPL-replaced enzyme is 90 to 95% inhibited by 10 mu CaC12. In order to investigate further the effect of DPL replacement, and subsequent solubilization with Triton, on the en-

4 Role of Protein and Lipid Motion in Ca +-ATPase Activity TABLE Effect of Triton X-100 and calcium on the rate of P, liberation at 0 C Rates of Pi liberation were measured at 0 C in a solution of 0.1 M KCl, 5 mu MgC12, 20 mm Tris/maleate, ph 7.0, 10 PM [-r- PI ATP. Unless specified, the free Ca2+ concentration was 10 PM. The final protein concentration was 0.2 mg/mi. A Triton X-lOO/protein weight ratio of 1.5 was used. Rate of P, liberation Additions None 10 mm CaClz Triton X-100 Triton X mm CaC12 I SR-ATPase DPL-ATPa& DPL-ATPase nmol mg- min the detergent might produce a minor change in the phosphorylation site. This change in the Ca2+ dependence, which is more marked with T&on-treated DPL-ATPase* (data not shown), would explain why the addition of Triton X-100 I- A zymatic behavior of the Ca +-ATPase, a detailed study of the formation of phosphoenzyme as a function of [Ca ] was carried out. To measure accurately the phosphoenzyme formation in the low [Ca ] range, it is essential to equilibrate the enzyme with the reaction solutions. While in SR or in SR- ATPase, incubation with Ca*+-free solutions results in an immediate inhibition of phosphoenzyme formation, in DPL- ATPase* an incubation time as long as 5 min in Ca2+-free medium is required to decrease the level of phosphoenzyme below 0.1 nmol/mg (Fig. 3). Preincubation in Ca +-free medium does not produce enzyme inactivation. If Ca2 is added to DPL-ATPase* after 15 min of incubation in the absence of Ca +, similar amounts of phosphoenzyme as those observed with the control (not preincubated) were formed without appreciable delay (Fig. 3). These results suggest that replacement with DPL results in a conformation of the enzyme in which, at low temperatures, a significant amount of Ca2+ remains tightly bound to DPL-ATPase*, requiring a long time for equilibration with the Ca +-free solution. This modification of the enzyme behavior by DPL seems to apply only to the removal of Ca2+. Once the tightly bound Ca2+ is removed, addition of Ca2+ to the reaction solution results in immediate phosphoenzyme formation (Fig. 3), indicating that the equilibration from Ca2+-free to Ca +-containing solutions is not affected by DPL replacement. The addition of Triton X-100 facilitates the removal of Ca2+ from DPL-ATPase*. In the absence of detergent, enough Ca2+ is still bound to the enzyme to allow significant phosphoenzyme formation after 1 min of incubation in Ca2+-free medium (Table II). However, in the presence of Triton a considerable reduction in phosphoenzyme formation was observed (Table II). The effect of Triton seems to be proportional to its concentration. The lowest phosphoenzyme formation after incubation for 1 min without Ca2+ was observed when Triton was added to DPL-ATPase* at the concentration at which it produces maximal stimulation of enzymatic activity (Table II). The complete profiles of phosphoenzyme formation as a function of [Ca ] for SR-ATPase, DPL-ATPase, and Tritonsolubilized DPL-ATPase show only minor differences among the three enzyme systems. Very little phosphoenzyme formation was observed below 10e7 M Ca2+ for all preparations (Fig. 4, A and B). The formation of phosphoenzyme increased between 10m7 and 10e5 M Ca2+ and remained constant up to 10e2 M Ca2+ for SR-ATPase and DPL-ATPase (Fig. 4A). In the case of the Triton-solubilized DPL-ATPase, a smooth increase in phosphoenzyme formation was observed from lo- to 5 x 10e4 M Ca2+; very little phosphoenzyme was formed below lo- M Ca2+ (Fig. 4B). This indicates that Triton-solubilized DPL-ATPase requires slightly higher Ca2+ concentrations for maximal phosphoenzyme formation, suggesting that )- 0 IO REACTION TIME, s FIG. 3. Time course of phosphoenzyme formation by DPL-ATPase* after variable preincubation times in Ca2+-free solutions. The reaction mixture contained 0.1 M KCI, 5 mm MgC12, 20 mm Tris/maleate, ph 6.5, 1.2 mm EGTA, 10 am [y- P]ATP, 0.2 mg/ml of protein, and Ca + as described below. The reaction was started by addition of [y- PJATP and was stopped at the times indicated in the abscissa by addition of 6.7% trichloroacetic acid. Phosphoenzyme formation was measured as described above. 0, phosphoenzyme formation in the presence of 1 mm CaCb ([Ca ] = 10 -, M), no preincubation. 0, phosphoenzyme formation in the absence of externally added Ca +. The reaction was started by addition of the enzyme (no preincubation in Ca +-free solution). 0, phosphoenzyme formation after 2 min of preincubation in Ca -free solution; A, phosphoenzyme formation after 5 min of preincubation in Ca +-free solution; A, phosphoenzyme formation upon addition of 1.0 mm CaC!b at time zero, after 15 min of preincubation in Ca +-free solution. TABLE Effect of Triton X-100 on phosphoenzyme formation by DPL- ATPase* after I-min incubation in Ca +-free solution Phosphoenzyme formation was measured at 0 C in a solution of 0.1 M KCl, 5 nuu MgC12, 20 mm Tris/maleate, ph 7.0, 10 PM [v- PI- ATP, 0.2 mg of protein/ml. Two different Ca2+ concentrations were used: 1Om5 M and a Ca +-free solution (less than lo- M) obtained by adding 1.2 mu EGTA to the medium described above. II Phosphoenzyme Additions 1 2 lo- M ca2+ CT+-fE!e nmol mg- Experiment 1 None Triton X-100 (1.5) Experiment 2 None Triton X-106 (0.6) Triton X-100 (1) n The numbers in parentheses represent the weight ratio of Triton X-100 to protein. 2/l

5 3.0 t IA Role of Protein and Lipid Motion in Ca -ATPase Activity B t PC P ca FIG. 4. A, phosphoenzyme formation by DPL-ATPase and SR- ATPase as a function of [Ca ] (pcu) at 0 C. The reaction mixture contained 0.1 M KCl, 5 mm MgCb, 20 mu Tris/maleate, ph 7.0, 10 PM [y-3zp]atp, and variable concentrations of Ca2+ and EGTA. The enzyme was equilibrated with the reaction mixture for 20 min at 0 C before starting the reaction with [Y-~ P]ATP. The final protein concentration was 0.2 mg/ml. The reaction was stopped by addition of 6.7% trichloroacetic acid after 30 s or 1 min of incubation. Phosphoenzyme was measured as described under Experimental Procedures. 0, DPL-ATPase; A, SR-ATPase. B, phosphoenzyme formation and rate of Pi liberation by DPL-ATPase treated with Triton X-100 as a function of [Ca For solubilization with Triton, the enzyme was incubated at a T&on/protein weight ratio of 1.5 for 5 min at 0 C. For the assay of phosphoenzyme formation and of P, liberation rates, the conditions were the same as described in A. 0, phosphoenzyme; 0, rate of P, liberation. results in a decrease of phosphoenzyme formation when assayed at micromolar Ca concentrations (cf: Fig. 2). The rate of Pi liberation for Triton-solubilized DPL-ATPase follows closely the pattern of phosphoenzyme formation up to 10e4 M Ca +, but is progressively inhibited from then on, reaching a very low value at lo- M Ca2+ (Fig. 4B). This same behavior has been reported for the membrane-bound ATPase enzyme in the absence of detergents (12, 20). Therefore, high Ca2+ concentrations have the same effect on the soluble form of the enzyme as on its membrane-bound form. Spin Label Experiments Lipid Motion-To monitor the fluidity of the lipid hydrocarbon chain region of the membrane, a stearic acid spin label was used (see Experimental Procedures ). The EPR spectra (at 4 C) of this probe incorporated into SR-ATPase and into DPL-ATPase* are shown in Fig. 5. The conventional spec- 1 trum of DPL-ATPase* (bottom left) indicates that the label is strongly immobilized, i.e. that the rotational correlation times characterizing the hydrocarbon chain motions are longer than 10e7 s and that the value of 2T li (64.8 G, Table III) is probably at or near the rigid limit, 2Tll (see Fig. 1) (16, 21). The conventional spectrum of SR-ATPase (top left) is similar, but the smaller value of 2T ll (60.2 G, Table III) indicates that the hydrocarbon phase is more fluid in this preparation. If we assume that 2Tl1, the rigid limit value of 2T ll for this spin label, is 64.8 G, then 60.2 G corresponds to an effective correlation time of 2 x loma s (from Fig. 1). Previous experiments comparing DPL-ATPase* with SR gave similar results (8). The difference in label motion between SR-ATPase and DPL-ATPase* is somewhat more apparent in the saturation transfer spectra (Fig. 5, right). Because of its sensitivity to slower motion, the saturation transfer spectrum of DPL-ATPase permits a quantitative estimate of the time range of the lipid motions, beyond the statement that the label is strongly immobilized : the L /L value is 0.82, indicating an effective rotational correlation time of about 10m4 s. The values of 2T ll and L /L for DPL-ATPase (Table III) are very similar to the values for DPL-ATPase*, indicating that the lipid phases of these preparations are very similar. In both cases, replacement of the native lipids with DPL decreases the lipid fluidity markedly, correlating with the inhibition of enzymatic activity (Table I). The addition of Triton X-100 to DPL-ATPase*, at a Triton/protein weight ratio of 1.5, results in a considerable increase in the motion of the stearic acid spin label (Fig. 6, broken line), as reflected in the large decrease in 22,, from 64.8 to 55.6 G (Table III). The result is a spectrum nearly identical with that obtained from the addition of Triton X- 100 to a sonicated suspension of DPL (2T ll = 55.0 G, Table FIG. 5. EPR spectra at 4 C of the stearic acid spin label incorporated into SR-ATPase (top) and DPL-ATPase* (bottom), at a ratio of one label/100 phospholipids. Left, conventional (VI) spectra. Rig/z& saturation transfer (V Z) spectra. The base-line is 100 G wide. TABLE III Separation of high and low field peaks in EPR spectra of SR- ATPase and DPL-ATPase* labeled with stearic acid spin label The enzyme preparations were labeled with 1 mol of stearic acid spin label per 100 mol of phospholipid. All spectra were recorded at 4 C. Triton X-100 was added at a Triton/protein weight ratio of 1.5. The number of observations (preparations) is in parentheses. The error involved in a single determination of 2T il is 0.1 G. Where more than one preparation was examined, the error given is the standard deviation. ~T II G SR-ATPase (3) SR-ATPase + 10 mm CaC (3) DPL-ATPase* (6) DPL-ATPase c 0.1 (1) DPL-ATPase* + Triton X f 0.9 (3) DPL-ATPase* + Triton X mm CaClz 55.6 f 0.9 (3) DPL + Triton X * 0.1 (1)

6 6884 Role of Protein and Lipid Motion in Ca2+-ATPase Activity and DPL-ATPase appear at least as large as those of SR. In addition, the saturation transfer spectra of SR-ATPase and SR are virtually identical. Therefore, in SR, SR-ATPase, and DPL-ATPase, any motions of the maleimide spin label that are detectable by saturation transfer spectroscopy (~2 c 10m3 s) are probably due to motions within the membrane, not to overall tumbling of membrane vesicles. Evidence against alternative b, the possibility of a loosely attached probe, is provided by the conventional spectra from MSL-SR-ATPase and MSL-DPL-ATPase in Fig. 7. The shapes of these spectra indicate that, regardless of the lipid composition or the concentrations of Triton and Ca2, the spin FIG. 6. Effect of Triton X-100 on the conventional EPR spectra of the stearic acid spin label in DPL-ATPase*. Solid line, no Triton; dashed line, 1.5 mg of Triton/mg of protein. The base-line of each spectrum is 100 G wide. III), at concentrations similar to those used in the enzyme preparations. These results suggest that after the addition of Triton, the lipid phase in DPL-ATPase* resembles a sonicated mixture of DPL and Triton. This large increase in lipid fluidity correlates with the large increase in enzyme activity. However, the addition of 10 mm CaClz to Triton-treated DPL-ATPase* or to SR-ATPase has little or no effect on the spectrum (see Table III), despite the strong inhibition of ATPase activity observed under similar conditions (see Table I). This finding indicates that the inhibitory effects of high [Ca ] are not mediated by a decrease in lipid fluidity. Protein Motion-In order to obtain more direct information about the effects of DPL, Triton X-100, and high [Ca ] on the physical state of the enzyme, we studied the rotational motion of a maleimide spin label attached covalently to the enzyme polypeptide (see Experimental Procedures ). Before we could use the spectra of this spin label to reliably determine the rotational mobility of the protein, we had to check two essential points (18). The first concerns the enzymatic behavior, which must not be drastically modified by labeling. Table IV shows the results of parallel experiments with DPL-ATPase and MSL-DPL- ATPase, in which the labeled preparation contains slightly more than 1 mol of label/lo5 g of protein. Although labeling causes a moderate reduction in both phosphoenzyme formation and Pi liberation rate, suggesting modification of essential -SH groups, the basic features of enzyme activity are unaffected by labeling; Pi liberation is stimulated by Triton and subsequently inhibited by high concentrations of Ca, and the inhibition of ATPase activity caused by DPL disappears at 37 C. Similarly, 1 mol of bound label/lo5 g of protein causes usually less than 50% inhibition of the activity of SR-ATPase. The second point concerns the nature of the rotational motion reported by the probe. It must correspond to the motion of the enzyme with respect to the membrane in which it is embedded and not to either (a) motion of the whole membrane vesicle or (6) motion of a loosely attached probe relative to the site of attachment on the enzyme. We have ruled out alternative a in the case of spin-labeled Ca -ATPase in SR vesicles at 4 C, by showing that the saturation transfer spectrum is almost unaffected by sedimenting the vesicles into a pellet, but that cross-linking the proteins with glutaraldehyde, without linking the vesicles together, slows the observed motion from a correlation time around 10F4 s to a correlation time of 10m3 s or longer. Electron micrographs of negatively stained preparations show that the vesicles of SR-ATPase ID. D. Thomas, and C. Hidalgo, (1978) Proc. h utl. Acad. Sci. l% S. A., in press. TABLE IV Enzymatic properties of DPL-ATPase and MSL-DPL-ATPase Rates of Pi liberation were measured as described in the legend to Table I. The final protein concentration was 0.2 mg/ml. A Triton/protein weight ratio of 1.5 was used. Rate of uhosuhate liberation 4oc 37 C nmol mg- mine DPL-ATPase x lo DPL-ATPase DPL-ATPase + Triton X Triton X mm CaCl2 MSL-DPL-ATPase MSL-DPL-ATPase + Triton X-100 MSL-DPL-ATPase + Triton X mivr CaCl? LOW co HIGH Ca A A SR-ATPass A A DPL-ATPcse I\ LOW Ca HIGH Ca x lo FIG. 7. EPR spectra at 4 C of MSL-ATPase and MSL-DPL-ATPase; effects of Triton and high [Ca ]. Parameters from these spectra are tabulated in Table V. A, conventional (VI) spectra. B, saturation transfer (V Z) spectra. As indicated in both A and B, spectra on the top row are from MSL-ATPase, those in the center row are from MSL-DPL-ATPase, and those in the bottom row are from Tritontreated MSL-DPL-ATPase (1.5 g of Triton X-100/g of protein). Low Ca (left column) corresponds to [Ca ] -lo- M. High Cu (right column) corresponds to the addition of 10 mm CaClz. The base-line of each spectrum is 100 G wide.

7 Role of Protein and Lipid Motion in Ca +-ATPase Activity labels are strongly immobilized (~2 > 10e7 s), indicating that there is little or no motion of the probe relative to its attachment site. Therefore, any changes in saturation transfer spectra are due to slow rotational motions (~2 > 1O-7 s) and probably correspond to large scale motions of the entire polypeptide or a large segment of it. Although the conventional spectra (Fig. 7A) of MSL-SR- ATPase and MSL-DPL-ATPase are not significantly different, the saturation transfer spectrum (Fig. 7B) of MSL-DPL- ATPase indicates much slower motion than in the case of MSL-SR-ATPase. The motion in SR-ATPase (Q = 6 x 1O-5 s, Table V), in the presence of endogenous SR lipids, is about the same as that observed in SR, whereas in DPL the observed motion ( X 10m4 s) is so slow that it is at or near the limit of resolution of the saturation transfer technique. Control experiments with partially delipidated SR- ATPase, containing (as does DPL-ATPase) 30 mol of phospholipid/l@ g of protein, indicate that the protein motion is only slightly slower than that observed with SR-ATPase, containing about 60 mol of phospholipid/l0 g of protein (data not shown). This indicates that the slow motion of DPL- ATPase is caused by DPL and not by its lower lipid content relative to SR-ATPase. The immobilization of the protein by DPL correlates with the concomitant reduction in lipid fluidity (Fig. 4 and Table III) and inhibition of enzyme activity (Table I, top line). The effect of Triton X-100 on the rotational motion of the enzyme in MSL-DPL-ATPase is shown in Fig. 7 and Table V. Although the conventional spectrum (Fig. 7A, bottom left) shows only a very slight decrease in 2T ll, possibly a qualitative indication of faster motion, the saturation transfer spectra (Fig. 7B, bottom left) indicate a dramatic increase in protein motion. The decrease in L /L from 1.13 to 0.51 (Table V) indicates a decrease in the effective correlation time by more than an order of magnitude. As discussed above, the enzyme is completely solubilized by Triton X-100. The membrane particles must be greatly reduced in size, to the point where their overall tumbling in solution affects the saturation transfer spectrum. The addition of 10 mm CaClz to Triton-treated MSL-DPL- ATPase does not significantly affect 2Tj, but it causes a substantial increase in L /L (Fig. 7B, bottom right, and Table V). Addition of lo- M CaCl2 produced a much smaller effect, and 10m4 M CaClz produced none (spectra not shown). Similarly, the addition of 10 mm CaClz to MSL-SR-ATPase results in a significant increase in L /L, corresponding to slower protein motion (Fig. 7B, top right, and Table V). We have observed a similar effect in SR membranes. Thus, both in Triton-treated DPL-ATPase and in SR-ATPase, (a) the protein rotational mobilities and enzyme activities are much greater than in DPL-ATPase, and (b) the addition of JO mm CaC12 produces a reduction in both protein motion and ATPase activity. Another means of relieving the enzyme inhibition caused by DPL, in addition to Triton treatment, is to raise the temperature above 29 C (7-9). We performed saturation transfer experiments as a function of temperature on MSL- DPL-ATPase and found that the effective correlation times below 29 C were all greater than 10e4 s, but were 10m4 s or less at temperatures above 29 C (Table VI). These results were obtained from a single preparation, so further studies should be carried out to clearly establish quantitatively the effect of temperature on the rotational motion of the CaZf-ATPase in different lipid environments. Furthermore, because the effects of temperature on saturation transfer experiments in model systems have not been studied in detail, care must be exercised in interpreting these data. Gel Chromatography One possible explanation for the effects of DPL, or high [Ca +], in decreasing the enzyme s rotational mobility would be the promotion of enzyme aggregation, which would result in larger, less mobile protein complexes. This model could apply to the effects of high [Ca ] on the soluble enzyme, as well as the effect of high [Ca ] or DPL on the membranebound enzyme. To test whether protein aggregation is responsible for the observed decrease in protein motion produced by addition of high [Ca ] to the soluble enzyme, SR-ATPase was treated with Triton X-100, plus or minus 10 mm CaC12, and fractionated on Bio-Gel A columns equilibrated with Triton, plus or minus 10 mm CaC12. The elution profile of the soluble enzyme under these conditions is shown in Fig. 8. In the absence of CaC12 the predominant protein species elutes at the position expected from a protein with an apparent molecular weight of 240,000, as determined from the elution positions of the two molecular weight standards used (cf Fig. 8). This is approximately the size of a dimer of the enzyme polypeptide. Lower amounts of protein appear at the positions expected for a monomer or trimer. Addition of 10 mm CaC12 causes an increase in the amount of monomer, at the expense of dimer and trimer (Fig. 8). No significant levels of higher molecular weight forms are observed. These results indicate that in the case of the Triton-solubilized SR-ATPase high [Ca2+] does not favor protein aggregation, but, in fact, shifts the size distribution to the monomeric form. TABLE EPR spectral parameters of maleimide spin-labeled enzyme; effects of DPL substitution, Triton, and high calcium concentration Triton was added in a Triton/protein weight ratio of 1.5. CaClz was added to 10 mu. Except for Rows 3 and 4, data correspond to spectra in Fig. 7. Each value is the average of several measurements from a single preparation; the uncertainties given indicate either the range of values observed or the estimated uncertainty of a single measurement, whichever was greater. The experiments were repeated with another preparation for Rows 1, 2, 5, 7, and 8; essentially the same results were obtained. Spectra were recorded at 4 C in a solution of 0.3 M sucrose, 20 mm Tris/maleate, ph 7.0, with the additions indi- cated above. SR-ATPase SR-ATPase + CaC12 SR-ATPase + Triton SR-ATPase + Triton + CaCh DPL-ATPase DPL-ATPase + CaC12 DPL-ATPase + Triton DPL-ATPase + Triton + CaCb Conventional WI) V Saturation transfer (WZ) ~T II L /L 72 G s z f c * r 0.1 TABLE VI c f t * x IO- 1.1 x 1o-4 4 x lo- 1.5 x lo- a x 1o-4 21 x lo- 2 x lo- 4 x lo- Effect of temperature on saturation transfer spectralparameters of maleimide spin-labeled DPL-ATPase Spectra were recorded at different temperatures in a solution of 0.3 M sucrose, 20 mm Tris/maleate, PH 7.0. The results were obtained from a single preparation. Temperature L /L 72 C x10-: x w x 1o x 1om x lo-

8 6886 Role of Protein and Lipid Motion in Ca +-ATPase Activity I t\ A * A I,A A FRACTION NUMBER FIG. 8. Chromatography of purified ATPase in Bio-Gel A 5m at 4 C. The solution loaded into the column (1 ml) contained 10 mg of nrotein dissolved in 15 mg of Triton X-100, 0.3 M sucrose, 20 mm %is/maleate, ph 7.0, plus TO) or minus (0) 10 mm CaCL The column was pre-eqmhbrated and eluted with a solution containing 0.1 M KCl, 20 mm Tris/maleate, ph 7.0, 15 mg/ml of Triton X-100, plus (0) or minus (0) 10 mu CaC12. Bovine serum albumin and carbonic anhydrase were used as molecular weight standards. DISCUSSION The enzymatic and spectral evidence presented in this work can be summarized as follows. (a) Replacement of the endogenous SR-ATPase phospholipids by DPL results in an enzyme preparation with a much less fluid lipid hydrocarbon phase at low temperatures. Whereas the effective correlation time of the enzyme associated with the SR lipids is less than 10m4 s, the rotational motion of DPL-ATPase is more restricted and has an effective correlation time -lo- s. In DPL-ATPase, the rate of Pi liberation is considerably lower than in SR-ATPase at 4 C, although their phosphoenzyme levels are basically identical. Only above 29 C, the temperature required to remove the inhibition of enzymatic activity produced by DPL, does the effective correlation time of DPL-ATPase decrease to less than lo- s. (b) DPL-ATPase is completely solubilized by Triton X The rotational motion of the protein becomes more than 10 times faster, and the motion of the stearic acid spin label incorporated into the lipid phase is also considerably enhanced. This solubilization is accompanied by a substantial increase in the rate of Pi liberation, reaching values similar to those found with SR-ATPase. SR-ATPase is also solubilized by Triton X-100 but its enzymatic activity remains unchanged. (c) The addition of high (10 mm) [Ca*+] to either SR- ATPase, Triton-solubilized SR-ATPase, or Triton-solubilized DPL-ATPase results in a strong inhibition of Pi liberation rates and in a decrease in the rotational mobility of the spinlabeled protein. It has little or no effect on the fluidity of the lipid hydrocarbon phase, as monitored by the stearic acid spin label. In the absence of Triton, high [CazC] has little or no effect on the already inhibited ATPase activity and protein motion of DPL-ATPase. In short, we observe a consistent correlation between ATPase activity and the rotational mobility of the protein in the sub-millisecond time range. We propose the following model to account for the effect of DPL on the behavior of the Ca *- ATPase enzyme. After replacement of the endogenous SR lipids by DPL, the protein is in a conformation that allows the active site to be readily phosphorylated by ATP, but the constraint imposed at low temperatures by rigid DPL molecules prevents the completion of the reaction cycle, i.e. phos- phoenzyme decomposition. There is a step of the reaction after phosphorylation that requires that the enzyme have a greater degree of rotational mobility than DPL permits. Our measurements indicate that this required rotational motion occurs in the sub-millisecond time range. This rotational motion does not imply the proposition that the enzyme functions as a rotary carrier. Without experiments on uniformly oriented planar membranes, we cannot determine the orientation of the motion relative to the plane of the membrane. However, since attachment to the enzyme of bulky molecules, such as antibodies against the ATPase (22) or against DNP cadaverine attached to the protein (23), results in little or no inhibition of ATPase activity and Ca2+ transport, it is unlikely that a 180 rotation about an axis in the plane of the membrane is required for enzyme function. Furthermore, in SR- ATPase the motions determined by EPR techniques are much faster than the motions required to account for the observed values of enzymatic activity, assuming a tightly coupled rotary carrier model. Our spectra are sensitive to rotational motion but not to translational motion. Thus, it is conceivable that the observed rotation is incidental to some other type of motion, e.g. a crucial shape change or a translational motion. However, it is very likely that phosphoenzyme decomposition requires some change in protein conformation or orientation (or both) which involves substantial movement of the enzyme (or of a large segment of it) relative to nearby lipids. Proposals that protein motion is required for enzyme activity have been made before for membrane-bound enzymes (24); we have presented here the first direct measurements of protein motion that support this type of model. The stimulation of Pi liberation produced at low temperatures by Triton X-100 can be explained, according to our model, as removal or fluidization of the rigid environment produced by the tightly packed DPL molecules around the enzyme polypeptide. Similarly, raising the temperature above 29 C increases the fluidity of the lipid environment of the enzyme (8). Either of these perturbations would allow the protein to increase its motion to the level required for normal functioning. The observed lack of effect of Triton on the Pi liberation rate in SR-ATPase at 4 C suggests that even at low temperature, the endogenous SR phospholipids already provide a sufficiently fluid environment around the enzyme to allow it to function at maximal efficiency. The effect of high [Ca2+] on the enzyme reaction has been extensively investigated for SR-ATPase (12, 20). It has been proposed that the Pi liberation rate is inhibited as a result of stabilization of the phosphoenzyme intermediate, and it has been shown that at O C, this stabilization takes place due to the binding of 2Ca2+/mol of phosphoenzyme, with an average affinity of lo3 M- (12). The present studies shed new light on this mechanism. In SR-ATPase, we have shown that the addition of high [Ca ] results in a decrease in the protein s rotational mobility. The most important finding is that similar CaZf-dependent changes occur in the enzymatic behavior and molecular motion of the detergent-solubilized enzyme, indicating that the stabilization by high [Ca ] does not require the existence of a phospholipid bilayer as a support for the enzyme. Furthermore, high [Ca ] has no substantial effect on the rotational mobility of the stearic acid spin label in SR- ATPase nor in Triton-solubilized DPL-ATPase. It is still conceivable that the Ca +-dependent stabilization process involves lipids that remain tightly bound to the protein, but this possibility is consistent with our results only if the fatty acid spin label is excluded from the lipid. protein complex. These results indicate that the stabilizing effect of high [Ca ], in contrast to that of DPL, is not mediated by the lipid phase, but is due to a change in the enzyme polypeptide itself that