Biophysics Rotational motion of the sarcoplasmic reticulum Ca 2 + - ATPase ( saturation transfer electron paramagnetic resonance / spin labels / membranes )

Using saturation transfer electron paramagnetic resonance, we have detected the rotational motion of a spin label rigidly attached to the sarcoplasmic reticulum Ca2 -ATPase (ATP phosphohydrolase, EC 3.6.1.3). At 40C, the spectrum indicates an effective rotational correlation time of 60 jsec, determined by comparison with reference spectra obtained from theoretical calculations and from experiments on model systems. This motion appears to correspond to rotation of the enzyme with respect to the membrane, because the motion persists when the membrane fragments are immobilized b sedimentation and the motion stops when the polypeptide cains, but not the membrane vesicles, are crosslinked by glutaraldehyde. The rotational mobility of the enzyme increases with increasing temperature, and this increase becomes more gradual when tie temperature exceeds 20'C; the same kind of temperature dependence has been observed previously for lipid fluidity and enzymatic activity. The dynamic nature of many processes occurring in biological membranes suggests strongly that molecular motions are extremely important aspects of membrane function. Therefore, direct measurements of motions of the lipid and protein components of membranes are essential to the understanding of the molecular details of membrane function. Translational and rotational motions of membrane lipids have been investigated extensively, as exemplified by the studies of spin-labeled lipids by McConnell and coworkers (1, 2). Some measurements have been made on the translational motion of membrane proteins (3, 4), but few measurements on their rotational motion have been reported (5, 6). Until recently, the magnetic resonance and optical spectroscopic techniques used to study the rotational motions of membrane lipids, usually in the nanosecond time range, were not sensitive to the much slower rotational motions that might be expected for membrane proteins. For example, the conventional electron paramagnetic resonance (EPR) technique, using nitroxide spin labels, can provide information only about motions characterized by rotational correlation times much less than 1 A.sec, whereas the rotational motion of a relatively small protein, rhodopsin, in a relatively fluid visual photoreceptor membrane is characterized by a rotational correlation time of the order of microseconds (5, 7). Saturation transfer EPR spectroscopy extends the time range of sensitivity to correlation times as long as 1 msec; this technique is therefore well-suited for studying the slow rotational motion of membrane proteins. The utility of the saturation transfer technique in the study of rotational motion in the microsecond to millisecond time range has been demonstrated in theoretical and model system studies (8, 9) and in studies on large-scale rotational dynamics in assemblies of muscle proteins (10). These and other applications of saturation transfer spectroscopy have been described in several recent reviews (11-14). Transport enzymes represent a particularly important kind of membrane protein in which rotational mobility may play a key role. We have focused our attention on the Ca2+-ATPase (ATP phosphohydrolase, EC 3.6.1.3) of sarcoplasmic reticulum (SR). Because this enzyme apparently requires a fluid lipid environment in order to function (15), it is likely that some degree of protein mobility, such as rotational mobility, is required for enzyme activity. In order to make possible the study of protein mobility as a function of enzymatic activity and lipid fluidity (16, 17) we have carried out a study of the rotational motion of the spin-labeled Ca2+-ATPase. We have attached a maleimide spin label selectively and rigidly to the enzyme in SR vesicles and used saturation transfer EPR to measure an effective rotational correlation time of 60 lisec at 40C, corresponding to rotation of the enzyme within the membrane. A preliminary account of some aspects of this work has appeared (17). EXPERIMENTAL PROCEDURES Membrane Preparations. Fragmented SR was prepared from rabbit white skeletal muscle as described (15). A purified Ca2+-ATPase preparation obtained by solubilization of SR with Triton X-100 (18), containing the endogenous SR lipids and a single 100,000-dalton polypeptide, will be designated SRATPase. A purified Ca2+-ATPase preparation in which the lipids in SR-ATPase have been replaced by dipalmitoyllecithin (DPL), as described (16), will be designated DPL-ATPase. Labeling with N-Ethylmaleimide. Unless otherwise indicated, the solution contained 0.3 M sucrose and 20 mM Tris maleate, pH 7.0 (SR buffer). SR, at a protein concentration of 10 mg/ml, was incubated with 1 mol of N-ethylmaleimide (MalNEt) per 105 g of protein for 1 hr at 00C and then was centrifuged at 100,000 X g for 1 hr to remove unreacted reagent. The pellet was rinsed and resuspended at a protein concentration of 10 mg/ml and then was incubated with maleimide spin label as described below. To determine the specificity of labeling under these conditions, the glycoprotein was separated from the ATPase by Triton X-100 solubilization of the SR vesicles (19). In order to determine the degree of MalNEt labeling, ['4C]MalNEt (New England Nuclear) was used as described (19). Protein concentrations were determined by the method of Lowry et al. (20), and ATPase activities were measured at 320C as described (8, 16). Spin-Labeling. A maleimide spin label derivative, N-(1oxyl-2,2,6,6-tetramethyl-4-piperidinyl)-maleimide (MSL), was used to label the enzyme polypeptide. A fresh solution of spin label in ethanol was diluted 1:10 with buffer, added to MalNEt-treated SR, SR-ATPase, or DPL-ATPase (final ethanol concentration, <1%) at a ratio of 1.25 mol of label per 105 g of Abbreviations: SR, sarcoplasmic reticulum; DPL, dipalmitoyllecithin; MalNEt, N-ethylmaleimide; EPR, electron paramagnetic resonance; MSL, maleimide spin label [i.e., N-(l-oxyl-2,2,6,6-tetramethyl-4piperidinyl)maleimide[. * Present address: Department of Structural Biology, Sherman Fairchild Center, Stanford University School of Medicine, Stanford, CA