ARCHIVES

OF BIOCHEMISTBY

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172, 12-19 (19%)

BIOPHYSICS

States of Myosin Subfragment-l Studied by Catalyzed Reduction of Bound Spin Label’

Ascorbate

HIROFUMI ONISH12 AND MANUEL F. MORALES3 Cardiovascular

Research Institute,

University

of California, San Francisco California 94143

Medical

Center, San Francisco,

Received May 2, 1975 The second-order rate constant, k, whereby ascorbate reduces spin label, N-(l-oxyl2,2,6,6-tetramethyl-4-piperidyl) iodoacetamide, bound to the fast-reacting (SH,) thiol groups of heavy meromyosin (HMM) has been compared with the k whereby ascorbate reduces free spin label in the same solvent. It is clear that the k of protein-bound spin label is primarily determined by conditions “on-board” subfragment-l (S-l), rather than by properties of the solvent. First, in saturating [STPI the k of HMM-bound spin label was much greater than the k of free spin label, and both k’s were independent of [KCll, from 0.05 to 1 M. Second, in the absence of ATP, or even in the presence of ADP, the k of HMM-bound spin label was less than the k of free spin label at 1 M KCl, and much more in a 0.05 M KCl. The organized structure of S-l is required for observing the change of K with ATP, because the change of k disappeared on denaturing HMM with either guanidine hydrochloride or urea. Measuring k can be a “probe” to specify HMM states. However, the parameter, k, is conceptually dissimilar to measuring peak heights on an EPR spectrum. Experimentally we have observed that when [KC11 is increased, while [MgATPl = 0, spectral peak height is constant, but k varies remarkably. At no [KC11 did excess F-actin affect k. Quantitative examination of metal contamination (e.g., Cu, Fe) in HMM showed that changes in the k of HMM-bound spin label cannot arise from changes in proximity to contaminating metal redox catalysts bound to HMM. An intramolecular participant in the reaction of ascorbate with bound nitroxyl half inhibits the Ca2+-ATPase of spin labeled HMM, so signal annihilation and ATPase activity are closely correlated in time. The rate of signal annihilation is unaffected by prior reaction of the “SH[ thiols with N-ethylmaleimide.

A small molecular “label” fixed to a microstructure can, under favorable conditions, 5eport” changes in its local environment as the microstructure undergoes various processes. But also, if there is a convenient third substance (a “quencher”) capable of annihilating the label signal, it is interesting to ask how readily quencher from the bathing solution can reach and * This research was supported by Grant CI 8 of the American Heart Association and Grant GB 24992X of the National Science Foundation. t Bay Area Heart Research Committee Fellow. Present address: Department of Chemistry, Science Faculty, Tokyo Institute of Technology, Tokyo, Japan. 3 Career Investigator of the American Heart Association.

4 Abbreviations used: HMM; heavy meromyosin; EPR, electron paramagnetic; TES, N-tris(hydroxylmethyl)methyl-2-aminoethane sulfonic acid; D’lT, dithiothreitol; MalNEt, N-ethylmaleimide; and EGTA; ethylene glycol-bis(P-aminoethyl ether)N, N’-tetraacetate. 12

Copyright All rights

0 1976 by Academic Press, Inc. of reproduction in any form reserved

annihilate the label signal, in different states of the microstructure. The information derived from quencher behavior is different from the information reported by the label itself, so the two experimental strategies usually complement one another. In this work we set out to study ascorbate quenching of an iodoacetamide-type spin label attached to each of the two most reactive thiols (SH,‘s) of heavy meromyosin (HMM),4 or of myosin. The behav-

MYOSIN

SPIN LABEL

ior of the label itself has been studied by us (l), by Stone (2), and, most thoroughly, by Seidel et al. (3, 4). Seidel (5) has also reported that dithiothreitol quenching of myosin-bound spin label is unaffected by myosin-actin binding. In our work, as in the work with the label itself, we have found that nucleotide binding has a strong effect, and actin binding has no effect, on ascorbate quenching. Additionally, we have found that KC1 has a strong effect on ascorbate quenching. Incidental to our quenching studies, we discovered an intramolecular reaction whereby a partic+~nt in ascorbate-reduction of nitroxyl inhibits the ATPase activity of HMM and of myosin. The protein group involved in this inhibition is not SH, (a thiol of myosin S-l whose reaction with N-ethylmaleimide extinguishes Ca2+ATPase). The existence of this inhibition does not invalidate the probing of myosin by measuring ascorbate quenching, since the inhibition does not occur until after quenching and apparently does not extend beyond the molecule whose nitroxyl is quenched. As usual, we have employed a spin label analog of iodoacetamide to effect low-degree labeling of the two reactive thiols on the S-l moieties of HMM. Typically, such HMM (18 PM) is exposed to 0.4 mM ascorbate, and th.e electron paramagnetic (EPR) spectrum is observed as a function of time. Because the EPR spectrum of the spin labels changes with nucleotide and KC1 variations, the decay of signal (measured as height from baseline, H) is observed at different field strengths under different conditions; this practice is justified by previous work (6) in which it was shown that the decay rate is the same at all frequencies. This previous work also established that under the conditions to be used here, ascorbate attack on bound or free nitroxyl is a pseudo first-order process, viz. H(t) is given by dHldt = k [ascorbatel H = K’H. Thus, k and k’ are second- and pseudo first-order rate constants, respectively; here we will report k as a parameter of probe state. METHODS Myosin was prepared from rabbit skeletal muscle as described by Tonomura et al. (7). The HMM was

13

isolated from a B-min trypsin digest of myosin cl:300 trypsin to myosin weight ratio, at pH 6.2) by the procedure of Lowey et al. (8) and concentrated by the procedure of Young et al. (9). G-actin was extracted from acetone powder of rabbit muscle at 0°C and purified by the procedure of Cohen (10). G-actin was polymerized with 0.1 M KCl, after removal of free ATP by treatment with anion exchange resin AG 1 x 4 E (Bio-Rad). Protein concentrations were determined by the Biuret reaction (11) calibrated by nitrogen determination or by ultraviolet absorption at 280 nm. The spin label, N-(lqxyl-2,2,6,6-tetramethyl-4-piperidinyl)iodoacetamide, was synthesized as described by McConnell and Hamilton (12); the corresponding amine derivative (tempoamine) was obtained from the Eastman Chemical Co. Heavy meromyosin (13.4 mg/ml) was incubated with 60 PM IAA spin label in a solvent containing 50 mM KC1 and 0.04 M Tris-HCl (pH 7.5) at 0°C for 60 min (with IAA, 80 pM IAA and 24 h were employed). After exposure, the HMM was dialyzed against 50 mM KC1 and 1 mM N-tris(hydroxylmethyl)methyl-2aminoethane sulfonic acid (TES) (pH 7.0) at 4°C for 2 days. Label (1.2-1.3 mol were found to bind covalently to a molecule of HMM, using the method of Stone (13). Spectra were recorded at room temperature with a Varian E-3 EPR spectrometer. The field strength was set at 3385 G, with a scan range of + 50 G. The modulation frequency was 100 kHz and the amplitude was 1 G. The microwave frequency was 9.536 GHz and the power was 50 mW. Labeled HMM was used at 5.5-6.5 mg/ml in the absence or presence of 0.5 mM ATP, or free label at 18-22 pM in 1 or 10 mM MgCl, and 0.05 or 0.08 ELM Tris-maleate (pH 6.5). Both 0.2 mg/ml pyruvate kinase and 0.20 mM phosphoenol pyruvate were used in an ATP-regenerating system. At zero time, the medium was made 0.4 mM in ascorbate. The peak heights were measured as a function of time at 3366 G (for free label), at 3355 G (for labeled HMM in the absence of ATP), and at 3362 G (for labeled HMM in the presence of ATP). Recording of the decay of spin signal was started 90 s after adding ascorbate to the system. To avoid microwave heating effects, cooling air was circulated through the cavity. A modulation amplitude of 4 G, instead of 1 G, was used in the case of bound labels to increase the signal-to-noise ratio. The concentration of Fe and Cu in HMM solution was measured with a Perkin Elmer Model 303 atomic absorption spectrophotometer equipped with a triple element (Fe-Ni-Cu) hollow cathode-lamp. ATPase activity was measured by determining the liberated Pi(t), using the conventional FiskeSubbarow method; at the concentrations used in this work, ascorbate ion did not perturb this assay. The sequence of measurement was as follows. After preincubation of labeled myosin or HMM with ascorbate in the proper solvent at room temperature for

14

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AND

various times, the system was rapidly diluted 250- to 309fold with incubation solution (0.6 M KCl, 10 mM pH 6.5, at 25°C). CaCl,, and 0.05 M Tris-maleate, ATPase was then started by adding 1 mM ATP. Ultrapure ammonium sulfate, ultrapure guanidine hydrochloride, and ultrapure urea were purchased from SchwarzJMann; ATP, ADP, pyruvate kinase, phosphoenol pyruvate and m-histidine were from Sigma Chemical Co.; L-glutamic acid was from Fisher Scientific Co.; and L-lysines were from both Pierce Chemical Co. and Calbiochem. Ascorbate was from a fresh bottle of Sigma reagent grade. In earlier work, no special precautions were taken to prevent ascorbate oxidation. In later work, wherever practical, solutions were preflushed with N, and were kept in specially sealed bottles. With these precautions, the solutions behaved as though they contained approximately 20% more ascorbate than when no precautions were taken.

MORALES

. hat progressive departure from linearity .s negligible for 6 5 pH 5 7, but becomes ncreasingly serious for pH 2 7, presumably due to auto-oxidation. On this ac.count all our work to be reported was at ,,H 5 7. Our comparisons between k for reducing HMM-bound spin label and k for reducing free spin label were made in the same solvent. The solvent components varied are: (i) 0.05 M I [KC11 I 1.0 M, a variation known to affect myosin solubility (151, the ATPase of modified myosin (16), and the EPR signal height ratio, Hz/H, (4); (ii) [MgATP] = 0, or 0.2 mM 5 [MgATPl 5 1.0 mM, a variation known to change the EPR signal height ratio, Hz/H, (4), and to impose a special complex, HMM**ADP * Pi, RESULTS on the ATPase site (17, 18); and (iii> The reduction of a protein-bound G!I- [MgADP] = 0, or 0.5 mM I [MgADPl 5 5.0 mM, a variation known to change slightly troxyl spin label is a pseudo first-order the EPR signal height ratio, Hz/H, (41, and process provided that ascorbate is sensibly constant throughout the experiment, so to impose the complex HMM.ADP on the ATPase site (19). we checked to make sure that auto-oxidaFigure 2A shows how MgATP affects the tion would not spoil this constancy under our conditions. At pH 7.0, using 0.4 mM rate of ascorbate attack on a spin label asc, we found that less than 2% is auto- bound to a special reactive thiol of HMM. label (0.35 min-’ oxidized in 15 min. The literature (14); sug- The k for HMM-bound mM-*) is elevated (to 0.85 min-’ rnM-‘) gests that the loss is more serious at higher pH. We studied the reduction of upon adding 0.5 mM ATP. Thirty minutes free spin label as H(t), and found (Fig. 1) after the addition of 1 mM ATP (at which time the ATP added is completely hydrolyzed), the k is the same level as k in the presence of 0.5-5 mM ADP (0.35 mine1 mM-l). It should be noted that due to the characteristics of the EPR spectra (Fig. 2B) different complexes are studied most conveniently by following the time-decay of maximum signal height, H, at different field strengths. Fig. 3A shows that varying [KC11 has no effect on the k for free iodoacetamide-type spin label; we do not show it here, but we also tested the fact that varying [MgATPl also has no effect on this k. Thus, it is obvious that the major changes in k shown for protein-bound labels are not due to the direct influence of FIG. 1. pH effect on ascorbate destruction of signal from free label. The label was used at 19 PM in 1 solvent features on these bound labels, but arise because solvent features influence M KCl, 10 mM MgCl,, 0.04 M buffer. Buffers were the protein, which in turn, influences the Tris-maleate at pH 6.0 (O), TES at pH 7.0 (A), and labels bound to it. Fig. 3A also shows that Tris-HCl at pH 7.5 (0) and 8.0 (V). At zero time, the at any [KCU, k of protein-bound label in medium was made 0.4 mM in ascorbate. The peak height at 3366 G was measured as a function of time. the presence of ATP is equal to or greater

MYOSIN SPIN LABEL

15

FIG. 2. EPR spectra and kinetics of ascorbate destruction of signal from HMM-bound label and free label. (A) Signal destruction, (Bl EPR spectra. Labeled HMM was used at 6 mg/ml, in the absence (A, solid line) or presence (Cl, broken line) of 0.5 mM ATP or free label at 18 FM (0) in 1 M KCl, 10 mM MgCl,, 1 mM EDTA and 0.08 M TES (pH 7.0).

A .

FIG. 3. KC1 effects on EPR spectra and kinetics of ascorbate destruction of signal from HMM-bound label and free label. (A) Rate constants of signal destruction. (B) EPR spectra. Labeled HMM was used at 5.5-6.5 mg/ml in the absence (A, solid lines) or presence (0, broken lines) of 0.5 mM ATP, and free label (0) at 22 &M in 10 mM MgCl, and 0.05-0.08 PM. Chelating agent (1 mM EDTA, 2 mM EGTA, or nothing) addition has little effect on k of ascorbate attack. [KC11 was 1 M (a), 0.3 M (b), or 0.05 M (c). Ascorbate solution was prepared with no special precautions (open symbols) in earlier work, and with precautions (filled symbols) in later work to prevent oxidation.

than k of free label; thus, it is also obvious sitive to both [KC11 and [MgATP]; in this that the changes in HMM brought about sense, observing the spectrum and measurby ATP binding to HMM do not consist in ing k are similar “probes” of HMM states. merely exposing the bound spin label to However, a spectrum relates fundamenthe bulk solvent. tally to a distribution of a population of Seidel and Gergely (4) have shown that systems among states (e.g., a very indirect features of the EPR spectrum are also sen- specification of enzyme topography);

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clearly they are conceptually dissimilar, and we should not expect them always to go hand in hand. For example, when [KC11 is increased, while [MgATPl = 0, spectral peak height, iY1, is constant (Fig. 3B), but k varies remarkably (Fig. 3A). The F-actin stimulates the ATPase of spin-labeled myosin (5). Whether it affects the EPR spectrum of myosin is controversial. Tokiwa (20) and Seidel (5) have reported an immobilization, but Stone (13) has reported no effect, suggesting that immobilization arises at labeled thiols other than the reactive (SH,) thiol. Table I shows that at high microwave power (which saturates the EPR signal of actolabeled HMM) excess actin, regardless of [KCII, has no effect on k for ascorbate attack. This result confirms Seidel’s similar result, viz., that F-actin has no effect on k for D’IT attack (5). Since the ascorbate attack on a nitroxyl label is a redox process, and since this process is sometimes faster on the HMM than in free solution, we had to consider the possibility of redox catalytic “site” on the HMM (for arguments given above they could not be merely in the solvent), sites existing there authentically (e.g., special amino acid residues), or created artiflcially (e.g., transition metal ions contaminating our HMM solutions). Accordingly, we sought experimental answers to certain pertinent, ancillary questions. Choosing arbitrarily a concentration of 25 PM for the metal ion, we measured k of free label in the presence of common metal ions that might contaminate HMM soluTABLE

DESTRUCTION BY ABCORBATE FROM HMM-BOUND LABELS (mghnl) 0 1.52 0 1.52

KCL

(M)

0.05 1.0

tions. We found that k for Co2+ and for Ni2+ are the same (0.56 min-’ mM-‘1 as for no addition, but for Fe3+ it is 0.62, and for Cu2+ it is 1.02 min-’ rnM-l. After these observations, our interest centered on Fe3+ and Cu2+. Two related questions arise: Is catalysis due solely to a protein-bound metal? Is the organized structure of S-l required for observing the change with ATP? These questions could be answered by first denaturing HMM with either 5 M ultrapure guanidine hydrochloride or 7.6 M ultrapure urea. After denaturation, k is the same for protein-bound label, for free label, and for protein-bound label in the presence of ATP. These results suggest that organized structure is necessary for the observed pattern of behavior of k, and that there are not, attached to the HMM, metal ions which alone can produce the special effects. We also were concerned with possible catalytic model compounds bearing some relation to amino acid residues that could exist on the S-l moieties of HMM. It was noted, for example, that after the aforementioned denaturations, free label k is elevated (to 0.79 min-’ mM-’ in 5 M guanidine hydrochloride and to 0.69 min-’ rnM-’ in 7.6 M urea) over k in the starting solvent, indicating some catalysis by these nitrogenous denaturating agents. We also found apparent catalysis by some amino acids. For instance, one sample of lysine (0.4 M) gave a free label k value in excess of 0.79 min-’ mM-‘, while another gave 1.44 min-1.5 Histidine (0.2 M) gave a 0.64 min-’ mM-1 and glycine (0.4 M) give 0.87 mine1 mM-1.

‘I

EFFECTOFF-ACTIN ON RATE CONSTANTS OF SIGNAL

F-actin

MORALES

k (mix’) 0.72 0.70 0.39 0.38

0.76 0.76

a Spin-labeled HMM was used at 4 mghnl, in 10 mM MgC&, 0.08 M Tris-maleate (pH 6.5), and 2 mM EGTA.

We found that complexes of Fe3+ and amino acids are better catalysts than either constituent alone. For example, 25 PM Fe3+ gives a k of 0.62 mini mM-l, and 0.4 M lysine gives a k of 0.79 min-’ m&r-l, but both together give a k of 1.82 mine1 mM-‘. Similarly, when 25 FM Fe3+ and 0.4 M glycine are mixed, the resulting k is 1.93 min-’ rnM-l. On the other hand, com6 These samples were found to be contaminated by 2.7 and 15.8 pM Fe, respectively, using direct atomic absorption spectrometry.

MYOSIN

SPIN LABEL

plexes of Cu2+ and amino acids seem to be ineffective; when Cu2+ and lysine are mixed, k is 0.79 mine1 rnM-’ (like lysine alone). Also, when Fe3+ is added to either the guanidine hydrochloride or urea (denaturing) solutions the k’s of the solutions are unchanged. The foregoing experiences with Fe3+ and Cu2+ made it important to know the contamination of these ions in our typical HMM solutions. By direct atomic absorption spectrometry we found 3.1 PM Cu and 0.7 PM Fe, i.e., the molar&y of Cu is approximately V12 and that of Fe is approximately l/50 the molarity of S-l moieties. Furthermore, even if these metals are added well in. excess (4 PM Cu2+, 1.5 and 3 PM in Fe3+) of their contamination level, k does not increase. We had to assure ourselves that ascorbate being used to attack the bound nitroxyl did not also have side reactions on myosin, in which case observations on how k depends on state might refer to “damaged” myosin. Our examination of this matter leads us to conclude that the myosin is damaged by ascorbate, but by a train of reactions subsequent to the attack on nitroxyl, so that we are reassured in our concern; however, these reactions seem sufficiently interesting to describe here. A “trivial” complication is that unless special precautions are taken, significant concentrations of dehydroascorbate form in stock ascorbate solutions. Drake et al. (21) have shown that this product of autooxidation can react with thiols; in the case of native myosin or HMM, reaction with the two especially reactive thiols (SH,‘s) occurs, and the Ca2+ ATPase of initially native enzyme is stimulated, particularly upon dilution. After uncovering this pitfall, we always used not only very fresh ascorbate solutions (as we had previously), but also only those kept in specially sealed containers, and, wherever possible, all systems studied were preflushed with N,. When such precautions were taken, ascorbate had no effect on the Ca2+ ATPase of native myosin. We are reporting some results obtained without these precautions

17

(e.g., some data of Fig. 3A), but only when tests had shown that the phenomena described were unaffected by the partial oxidation of the ascorbate. Independently of purification, ascorbate inhibits the Ca2+ ATPase of spin-labeled myosin or HMM, rapidly at 2 mM, slowly at 0.4 mM, but the same nonzero extent (Fig. 4). Different degrees of labeling of course produce different Ca2+ ATPase activities, but the asymptotic percentage of inhibition (about 40%) by ascorbate is independent of labeling degree. Since native enzyme is unaffected by ascorbate, but nitroxyl-labeled enzyme is affected, we believe that the inhibitory agent is a product of the reaction of ascorbate and nitroxyl. In an effort to define this product we investigated whether it is diffusible from one enzyme molecule to another. The reactions of IAA and IAA spin label with myosin are so similar as to indicate that both labels react with the same (SH,) thiol. Ascorbate attacks “tempoamine” as well as spin-label, free in solution. However, when IAA-labeled myosin was incubated with tempoamine and ascorbate the Ca2+ ATPase of the myosin was not affected, strongly suggesting that all par-

FIG. 4. Inhibition of Cae+-ATPase of labeled myosin. After preincubation of 9.8 mg/ml labeled myosin with ascorbate in 0.6 M KCl, 10 mM CaCl,, and 0.1 M L-histidine-HCl (pH 6.5) at room temperature for various times. Myosin solutions were diluted 300-fold for ATPase measurements. Concentration of ascorbate was 0.4 mM (0, 0) or 2 mM (A). Degree of labeling was 1.17 mol (0) or 1.93 mol (0, wt) per myosin molecule.

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ticipants in the inhibition reaction are confined to the same molecule, i.e., the participant may be an altered group of myosin. If the ATPase inhibition is directly linked to the attack of ascorbate on nitroxyl, then signal height should disappear at the same rate as ATPase disappears. That this is so is shown in Fig. 5 for two different concentrations of KCl. We suspected that the myosin sites at which inhibition occurs are the so-called SH, groups, whose ligation inhibits ATPase, so we studied spinlabeled myosin whose ATPase had been virtually destroyed by reaction with N-ethylmaleimide (MalNEt). Capitalizing on the above correlation between signal decay and inhibition, we incubated the MalNEt-treated myosin with ascorbate. However, signal still decayed, almost as in untreated myosin, suggesting that our suspicion was incorrect and that the ascorbate inhibition does not occur at the SH, group.

FIG. 5. Inhibition of Ca2+-ATPase and ascorbate destruction of signal from HMM-bound label. Labeled HMM (1.38 mol/HMM molecule) at 5.5 mg/ml was incubated with 0.25 mM ascorbate in 0.05 M (A) or 1 M (0) KCl, 1 mM MgCl,, 0.05 M Tris-maleate (pH 6.5) at room temperature for various times. Incubation was terminated by a 250-fold dilution of HMM solution prior to ATPase measurement. For ascorbate destruction of signal, the medium used was the same as the incubation medium, except that it contained an ATP regenerating system. Concentration of KC1 was 0.05 M (+) or 1 M (X).

MORALES DISCUSSION

Our results show: (i) When [KC11 is increased, ascorbate attacks the bound spin label with decreased effectiveness. (ii) Generally speaking, at any fixed [KCll, if MgATP is added, the effectiveness of the ascorbate attack is increased, always to the same maximum. The increase itself depends on the fixed [KC11 and there is the suggestion in our data that ATP-addition restores the effective attack condition that KCl-addition progressively eliminates. (iii) The changes in label vulnerability to ascorbate are not simply a matter of changing exposure to bulk solvent, they are not exhibited by unbound label, and therefore must consist in shifting the label from one environment in the protein to another environment in the protein. (iv) Whatever the environmental changes in (i)-(iii) are, Factin binding has no effect upon them. If as one adds KC1 in the range, 0.05-1.0 M, there is a progressive change around the spin label (expressed as a decreasing vulnerability to ascorbate), one must consider that the change may be correlated with the progressive decrease in H,IH, ratio (4) and in ATPase (161, but these considerations must be left to future research. In what regard do the label environments that we have postulated differ from one another? A natural speculation is that KC1 addition withdraws the label from a redox catalytic site and ATP restores proximity. But what is the nature of a “redox catalytic site?” Since we have found complexes of metals and amino acids to be better catalysts than either constituent alone, we could speculate that the site is such a complex. The simplest version of this hypothesis, however, that the metal is bound to a particular residue (or residues) forming a catalytic site (or sites), is contradicted by our data, for analysis shows that in our systems there are far fewer common redox metals than S-l moieties; nevertheless, all of the signal is destroyed. Under such circumstances, we should observe more than one rate (h) of ascorbate attack, but in fact we observe only one. Also these metals cannot simply be rapidly exchanging among high affinity sites because if we add metals to the system, k is unaffected.

MYOSIN Thus,

We

are

driven

b

think

that

if

19

SPIN LABEL a

redox catalytic site exists it is constituted from myosin residue (or residues). The best accounting for our various observations can be made if we assume that on the myosin surface there exists a group that accepts electrons from ascorbate and donates them to nitroxyl, but whose completely reduced form is incompatible with active ATPase. In this hypothesis catalyzed reduction would be strictly correlated with ATPase destruction, and effectiveness of the catalysis (L?value) would be modulated ‘by environmental distortions, e.g., by how proximal or well-oriented is the myosin group to the nitroxyl. Thus, we would say that variations in [KCl], or imposition of steady-state ATPase (i.e., imposition of the species now commonly called myosin ’ **ADP, Pi), produces environmental distortions that are sensed by our measurement. To account for additional observations that we have made, the hypothesis has to be expanded. Because the k for myosin-bound label can be less (at high [KCll), as well as greater, than the k for free label, we have to think that at least some of our phenomena occur in a crevice shielded from solvent, an idea made quite plausible by others (Haugland, R.P., submitted for publication). Finally, we have to leave two important questions largely unanswered: Why is there a limit to the ATPase inhibition, but not to signal destruction? What is the myosin group involved in the redox catalysis? The last question remains unanswered, but we can say that the group is not the reactive thiol (SH,) itself (because the label is attached there), and that it is not the thiol called SH, (because attaching MalNEt to SH, does not affect signal destruction rate). d4CKNOWLEDGMENTS We are indebted, for important advice, to Prof. Y. Tonomura, Dr. Roger Cooke, and Dr. Robert Mendel-

son, and, for kindly supplying us with the sample of IAA spin label, to Dr. Richard Haugland. We also thank Mrs. Susan L. Putnam for skillful assistance. Professor J. Botts kindly made her atomic absorption spectrometer available to us. REFERENCES 1. QIJINLIVAN, J., MCCONNELL, H. M., STOWRING, L., COOKE, R., AND MORALES, M. F. (1969) Biochemistry 8, 3644. 2. STONE, D. B. (1970) Arch. Biochem. Biophys. 141, 378. 3. SEIDEL, J. C., CHOPEK, M., AND GERGELY, J. (1970) Biochemistry 9, 3265. 4. SEIDEL, J. C., AND GERGELY, J. (1973) Arch. Biochem. Biop ys. 158,853. 5. SEIDEL, J. C. (1973) Arch. Biochem. Biophys. 157, 588. 6. TONOMURA, Y., AND MORALES, M. F. (1974) Proc. Nat. Acad. Sci. USA ‘71, 3687. 7. TONOMURA, Y., APPEL, P., AND MORALES, M. F. (1966) Biochemistry 5, 515. 8. L~WEY, S., SLAYTER, H. S., WEEDS, A. G., AND BAKER, H. (1969) J. Mol. Biol. 42, 1. 9. YOUNG, D. M., HIMMELFARB, S., AND HARRINGTON, W. F. (1965) J. Biol. Chem. 240, 2428. 10. COHEN, L. B. (1966) Arch. Biochem. Biophys. 117, 289. 11. GORNALL, A. G., BARDMILL, C. J., AND DAVID, M. M. (1949) J. Biol. Chem. 11’7, 751. 12. MCCONNELL, H. M., AND HAMILTON, C. L. (1968)

Proc. Nat. Acad. Sci. USA 60, 776. 13. STONE, D. B. (1973) Biochemistry 12, 3672. 14. WEISSBERGER, A., LUVALLE, J. E., AND THOMAS, D. S., JR. (1943) J. Amer. Chem. Sot. 65,1934. 15. BRAHMS, J., AND BREZNER, J. (1961) Arch. Bio-

them. Biophys. 95,219. 16. WARREN, J. C., STOWRING, L., AND MORALES, M. F. (1966) J. Biol. Chem. 241, 309. 17. VINIEGRA-GONZALEZ, G., AND MORALES, M. F. (1972) Bioenergetics 3, 55. 18. TRENTHAM, D. R., BARDSLEY, R. C., ECCLESTON, J. F., AND WEEDS, A. G. (1972) Biochem. J. 126, 635. 19. LOWEY, S., AND LUCK, S. M. (1969) Biochemistry 8, 3195. 20. TOKIWA, T. (1971) Biochem. Biophys. Res. Com-

mun. 44,471. 21. DRAKE, B. B., SMYTHE, C. V., AND KING, C. G. (1942) J. Biol. Chem. 143, 89.