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. 1997 Sep 2;94(18):9643–9647. doi: 10.1073/pnas.94.18.9643

Independent mobility of catalytic and regulatory domains of myosin heads

Bishow Adhikari *, Kalman Hideg †, Piotr G Fajer *,‡
PMCID: PMC23242  PMID: 9275176

Abstract

The recent determination of the myosin head atomic structure has led to a new model of muscle contraction, according to which mechanical torque is generated in the catalytic domain and amplified by the lever arm made of the regulatory domain [Fisher, A. J., Smith, C. A., Thoden, J., Smith, R., Sutoh, K., Holden, H. M. & Rayment, I. (1995) Biochemistry 34, 8960–8972]. A crucial aspect of this model is the ability of the regulatory domain to move independently of the catalytic domain. Saturation transfer–EPR measurements of mobility of these two domains in myosin filaments give strong support for this notion. The catalytic domain of the myosin head was labeled at Cys-707 with indane dione spin label; the regulatory domain was labeled at the single cysteine residue of the essential light chain and exchanged into myosin. The mobility of the regulatory domain in myosin filaments was characterized by an effective rotational correlation time (τR) between 24 and 48 μs. In contrast, the mobility of the catalytic domain was found to be τR = 5–9 μs. This difference in mobility between the two domains existed only in the filament form of myosin. In the monomeric form, or when bound to actin, the mobility of the two domains in myosin was indistinguishable, with τR = 1–4 μs and >1,000 μs, respectively. Therefore, the observed difference in filaments cannot be ascribed to differences in local conformations of the spin-labeled sites. The most straightforward interpretation suggests a flexible hinge between the two domains, which would have to stiffen before force could be generated.

Keywords: muscle, light chain, spectroscopy, EPR, saturation transfer–EPR

The myosin head (subfragment 1, S1) plays a central role during muscle contraction. It is thought that during the ATPase cycle S1 interacts with actin and undergoes a series of specific changes in its conformation, resulting in a directional strain on the actin filaments. The nature of these conformational changes and the mechanism by which they produce the directed force is still unclear (1, 2), although the original model of Huxley and Simmons (3) provides a valid framework. The determination of the atomic structures of S1, actin, and the low-resolution structure of the acto-S1 complex has led to a hypothesis of a structural model, in which helix movement is generated by the hydrolysis of ATP and the formation of a stereospecific acto-S1 complex generates torque within the catalytic domain, which is then amplified by the regulatory domain acting as a lever arm (4, 5).

Rayment’s model was preceded by the observation of shape changes in the myosin heads between various chemical states of myosin. A decrease in the hydrodynamic size of S1 upon MgADP binding and upon ATP hydrolysis was observed by Highsmith and Eden (6) and by Wakabayashi et al. (7, 8). More recently, low-resolution electron microscopy reconstitution of S1-decorated actin filaments and EPR studies have indicated a rotation of the regulatory domain with respect to the catalytic domain (9–11). In these cases, the rotation was observed at the distal end of the regulatory domain on the binding of ADP only in smooth muscle S1 or brush border S1. Studies were not carried out in skeletal muscle; thus, the question remained whether there is a similar rotation in skeletal muscle and on the location of the possible hinge. The atomic structure of myosin S1 strongly suggests the interface between the catalytic domain and the essential light chain is a putative hinge (12).

We spin labeled two cysteine residues, 31Å apart, on either side of the putative hinge—Cys-707 in the catalytic domain and Cys-177 of the essential light chain in the regulatory domain—and compared their mobilities in filaments made of skeletal myosin. Saturation transfer–EPR (ST-EPR) spectroscopy was utilized in probing domain movements because ST-EPR is uniquely sensitive to the microsecond motions expected for the myosin head. We found that the mobility of the catalytic domain is three to five times higher than that of the regulatory domain. This finding implies that the connection between the two domains is flexible, lending further support to models in which internal shape changes within the myosin head lead to force generation (4, 13, 14).

METHODS

Protein Purification.

All preparative procedures were carried out at 4°C. Rabbit myosin was extracted with Guba-Straub solution (0.3 M KCl, 0.05 M K2HPO4, 0.1 M KH2PO4) and purified by repeated polymerization–depolymerization cycles in low and high ionic strength (μ) buffers as described by Margossian and Lowey (15). Purified myosin was either directly used or stored in 50% glycerol at −20°C. A mixture of essential light chains (ELCs) containing A1 and A2 isoforms was prepared from purified myosin by removal of myosin heavy chain from the light chains by guanidine-HCl denaturation followed by ethanol precipitation (16), except for an additional repeat of the procedure for the regulatory light chain removal. The procedure yielded an ≈95% pure essential light chain mixture upon examination by 12% SDS/PAGE gels. Purified ELC (1–2 mg/ml) was stored at −70°C in 5 mM phosphate buffer, 1 mM NaN3, and 0.1 mM DTT at pH 7.0 (storage buffer). Myosin rods were prepared by chymotryptic digestion of myosin to remove S1 and soluble heavy meromyosin followed by removal of residual myosin by solubilization and denaturation in high salt and 95% ethanol (15). Actin was prepared from acetone powder by extracting in a low μ buffer followed by cycles of polymerization and depolymerization induced by μ changes according to Pardee and Spudich (17).

Spin Labeling.

Purified ELC mixture in storage buffer was dialyzed in 40 mM KCl, 0.2 mM EDTA, 1 mM NaN3, and 5 mM phosphate buffer at pH 7.0 and labeled with a 5-fold molar excess of spin label for 5–12 hr, followed by dialysis in the same buffer. A labeling ratio of 80–100% was achieved, as determined by spin quantitation using EPR and protein concentration determination using the BCA protein assay (Pierce). The catalytic domain of S1 was labeled with indane dione spin label (InVSL) according to Roopnarine et al. (18). Myosin rods were labeled in 0.6 M KCl, 50 mM Mops, pH 7.0, with a 5-fold excess of InVSL overnight, followed by dialysis in the same buffer to remove the excess spin label. To form filaments, labeled myosin and rods were dialyzed in 40 mM KCl, 10 mM Mops, pH 7.0, centrifuged at 10,000 × g for 10 min., and loaded into 50-μl fused silica capillary tubes (Wilmad, Buena, NY) for EPR experiments.

Spin-labeled ELCs (InVSL-ELCs) were immobilized on preactivated diisothiocyanate (DITC) glass beads (Sigma) by incubation of the InVSL-ELCs in buffer-prewashed DITC beads overnight, at an equimolar concentration of protein and binding sites on the beads, followed by removal of free light chains by several washes in the same buffer.

The spin-labeled light chains were exchanged into myosin according to the method of Wagner (16). Typically, 40–80% exchange of labeled ELC per head was achieved, as determined by spin quantitation and the BCA assay. The exchange of labeled ELC into myosin did not significantly (<10%) alter the ATPase activity (Table 1).

Table 1.

ATPase activities* of InVSL-ELC-exchanged myosin and the native myosin

Condition Native InVSL-ELC exchanged
Physiological ionic strength† 0.016  ± 0.002 0.016  ± 0.002
Low ionic strength† 0.026  ± 0.006 0.022  ± 0.006
Actomyosin in low ionic strength 0.5  ± 0.03 0.4  ± 0.02
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*

ATPase activities are reported as μmol of Pi/min per mg of myosin. The actomyosin activity was carried out with a 7.5–10 molar excess of actin over myosin. All experiments were carried out at 25°C. 

†

The physiological ionic strength buffer consisted of 130 mM potassium propionate, 20 mM Mops, 2 mM MgCl2, 2 mM EGTA, 1 mM NaN3, pH 7.0. The low-ionic-strength buffer was 40 mM salt and 10 mM Mops, pH 7.0. 

EPR Experiments.

EPR and ST-EPR experiments were carried out as in Adhikari and Fajer (19), using a TE102 cavity with microwave field strength (H1) of 0.03–0.08 G and microwave modulation amplitude (Hm) of 2 G for EPR and H1 = 0.25 G and Hm = 5 G for ST-EPR. The modulation frequency was 50 kHz. All experiments were carried out at 4°C. The ST-EPR spectra were analyzed by comparison of the experimental line shapes with those obtained from InVSL-labeled hemoglobin tumbling in media of known viscosity [87.55% wt/wt, glycerol/water mixture, −30° to ≈20°C (20)]. Conventional EPR spectra were analyzed as described by Freed (21), with rotational correlation time given by τR = a(1 − 2Teff/2Tmax)b, where 2Tmax is the rigid limit for a particular spin-labeled protein and 2Teff is the splitting between the outermost peaks, a = 5.4 × 10−10 s, and b = −1.36 for Brownian motion.

RESULTS

Spin-Labeling ELCs.

Three spin labels, InVSL, iodoacetemide (IASL), and maleimide (MSL) nitroxides, were tested in an attempt to find a rigidly attached probe on ELCs that will accurately report domain or global motions rather than the independent motion of the spin label relative to the protein. In solution, the spectra of labeled ELCs indicated rapid nanosecond motions in the following order of increasing rotational correlation times: InVSL-ELC, 1.2 ns; IASL-ELC, 1.3 ns; and MSL-ELC, 1.5 ns (Fig. 1A, IASL and MSL spectra not shown). These mobilities are faster than the τR = 8 ns predicted from the Stokes–Einstein equation.§ Faster than expected mobilities of spin-labeled protein imply librational motion with respect to the isolated light chain or very fast mobility of the spin-labeled domain.

Figure 1.

Figure 1

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Rigidity of InVSL attached to Cys-177 of LC1. Conventional EPR spectra of InVSL-labeled ELCs in solution (A) and after exchange into myosin filaments formed by dialysis (B). The narrow effective splitting (2Teff) in the solution spectrum, which indicates rapid nanosecond motions, becomes broad and large in the myosin filament spectrum, thus indicating the absence of fast nanosecond motions. Immobilization of the labeled ELCs on DITC-coated glass beads also resulted in an EPR spectrum (C) with a large 2Teff, indicating rigid attachment of the label. Likewise, the ST-EPR spectrum (D) from the immobilized sample with the L"/L line-height ratio of 1.5 is characteristic of an effective rotational correlation time (τR) in the millisecond time scale. The samples were in 40 mM KCl, 10 mM Mops, pH 7.0, and 23°C.

Fortunately, this fast mobility was reduced when the InVSL-ELC was exchanged into myosin (Fig. 1B) or into S1 (spectra not shown). The largest immobilization was observed with InVSL. Compared with the sharp spectrum in solution, the spectrum from myosin filaments was broad, with the hyperfine splitting, 2Teff = 71.0 ± 0.21 G, which is the rigid limit for this label. The spectra of IASL and MSL were only slightly broadened (not shown), implying considerable librational motions relative to the protein, τR = 1.9 ns and τR = 1.6 ns, respectively, rendering them unsuitable for the study of regulatory domain dynamics.

To further examine the rigidity of InVSL with respect to ELC, the spin-labeled protein was immobilized on DITC-coated glass beads. Both the conventional and ST-EPR spectra were at the rigid limit (Fig. 1 C and D, respectively). The splitting of the EPR spectrum is 70.7 ± 0.22 G and the diagnostic line height ratio, L"/L, of the ST-EPR spectrum is 1.5. Thus, the InVSL label is immobilized both on the nanosecond time scale of EPR and on the millisecond time scale of ST-EPR experiments, and therefore can be used to study the dynamics of the regulatory domain. The InVSL was previously shown to be rigidly attached to Cys-707 of the catalytic domain using the same criteria (18).

Mobility in Myosin Filaments.

The spectra of the myosin filaments formed at low ionic strength (μ = 45 mM) with spin labels in the catalytic domain (Cys-707), regulatory domain (Cys-177), and myosin rod are shown in Fig. 2. A significant difference in mobility is apparent between the spectra of the catalytic and regulatory domains. The higher intensity of the diagnostic feature L" in the spectrum of the regulatory domain reflects the slower mobility of this domain as compared with the catalytic domain spectrum. The effective τR, calculated using the L"/L line height ratio, is 36 ± 12 μs for the regulatory domain and 6.9 ± 1.5 μs for the catalytic domain (Table 2). The latter value is in agreement with a previous ST-EPR estimate (22) or from time-resolved optical anisotropy (23, 24). The mobility of both domains of the myosin head was higher than the mobility of myosin rods, τR = 165 ± 50 μs, consistent with a hinge at the head-rod junction, enabling myosin head movement relative to the myosin filament backbone (Fig. 2). The flexibility of that particular hinge is modulated by ionic strength, pH, phosphorylation of regulatory light chain, or presence of MgATP (25).

Figure 2.

Figure 2

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ST-EPR spectra comparing the mobility of the regulatory domain (Top), the catalytic domain (Middle), and the rod (Bottom) in synthetic filaments prepared by dialysis at μ = 45 mM (see Methods). The spectrum of the regulatory domain indicates lower mobility for this domain than that of the catalytic domain. The samples were in 40 mM KCl, 10 mM Mops, pH 7.0, and 4°C.

Table 2.

Comparison of effective correlation times of the two head domains

Condition Effective τR, μs
Regulatory domain Catalytic domain
Myosin monomers* 2.6  ± 0.6 (3) 2.7  ± 1.6 (3)
Myosin filaments† 36  ± 12 (6) 6.9  ± 1.5 (14)
Actomyosin* 1,000–2,000 (3) 1,000–3,000 (2)
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Effective correlation time is defined as that of InVSL-hemoglobin displaying the same L"/L line-height ratio. The numbers enclosed in brackets indicate the number of measurements. 

*

In 0.6 M KCl, 20 mM Mops, pH 7.0 at 4°C. 

†

In 40 mM KCl, 10 mM Mops, pH 7.0 at 4°C. 

Mobility in Myosin Monomers and Actomyosin.

It is possible that the difference in mobility between the two domains arises from the interaction of the label with its immediate environment of various amino acid residues surrounding the labeled sites. For example, the different orientation of the spin label principal axes with respect to the diffusion axis might result in a different ST-EPR line shape for the same motional mode (26). To exclude this possibility, the mobility of the two domains was measured in monomeric myosin and in actomyosin. If the line-shape difference was due to the local environment, then we would expect the difference line shape to persist in actomyosin or monomeric myosin. Fig. 3 shows a representative set of spectra for the two domains in monomeric myosin.

Figure 3.

Figure 3

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ST-EPR spectra of the regulatory (Upper) and the catalytic domains (Lower) of myosin heads in monomeric myosin. The samples were in 0.6 M KCl, 20 mM Mops, pH 7.0, and 4°C.

Both spectra indicate faster mobility than was observed for the filament form of myosin, consistent with greater rotational flexibility in the monomeric form than in the filament form. Importantly, no significant difference was observed in the spectra between the two domains. The correlation times were 2.6 ± 0.6 μs for the regulatory domain and 2.7 ± 1.6 μs for the catalytic domain (Table 2). Likewise, when the myosin head is immobilized by actin, the mobility of the two domains is equal; τr = 1–2 ms (Fig. 4).

Figure 4.

Figure 4

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ST-EPR spectra of the regulatory (Upper) and the catalytic (Lower) domains of myosin in the presence of actin filaments. The samples were in 0.6 M KCl, 20 mM Mops, pH 7.0, and 4°C.

The absence of any mobility differences between the catalytic and regulatory domains in the two motional extremes—monomeric form and actomyosin—strongly suggests that neither differences in the local environment of the label sites nor the different anisotropy of label motion at the two sites contributes toward the difference observed in the myosin filaments (26).

DISCUSSION

The present study demonstrates that the regulatory and catalytic domains of the myosin head are capable of independent movement, indicating that the connection between the two domains is flexible. The difference in the measured effective τR from the two domains may be due to differences in rates of motion or amplitude, or a combination of both (27). The possibility of changing probe environment to account for this difference was ruled out by identical τR in the two domains in actomyosin and monomeric myosin, suggesting that the local environment of the probed sites in catalytic and regulatory domains are the same. The finding that τR of the two domains varies as expected in monomers, in filaments, and when bound to actin, strengthens the validity of using rotational correlation time for probing the relative movement of myosin head domains in various conformations.

The mobility of the two domains might be modeled using the formalism developed by Broersma (28) as modified by Hagerman and Zimm (29). To model the relative motions of the regulatory and catalytic domains, let us assume that the mobility of the regulatory domain equals that of the entire head, which is approximated by a semi-rigid cylinder with 5-nm diameter and 17-nm length and is attached to the thick filament shaft by a universal joint (S1/S2 junction). The catalytic domain might be modeled as a rigid cylinder half the length of the head (8.5 nm) and attached to the regulatory domain by a second universal joint (catalytic-regulatory-domains hinge). The predicted difference in the diffusion coefficients between the regulatory and catalytic domains is then 5.5-fold, while the observed values differ by 5.3. This good agreement between the observed and predicted values supports the validity of the original assumption of low motional restrictions placed on the mobility of the catalytic domain by the regulatory domain.

Myosin heads in filaments possess considerable freedom of movement even though the rod and part of S2 form the rigid filament backbone, as established previously for the catalytic domain (22–24) and in this work for the regulatory domain. The larger increase in τR of the regulatory domain compared with the catalytic domain upon filament formation indicates that the hinge at S1/S2 may not be completely flexible, perhaps due to interactions between this domain and the filament backbone. Indeed, phosphorylation of regulatory light chain, which results in the movement of the head away from the filament backbone and an increase of regulatory domain mobility, suggests that these interactions may be related to force modulation (25, 30).

The most recent model of muscle contraction, based on the crystal structure of the myosin head, postulates a domain displacement as necessary for the generation of the power stroke (4). A comparison of the crystal structures of the catalytic domain from Dictostelium discoideum complexed with beryllium fluoride and aluminum fluoride, thought to mimic the ATP bound state and the transition state for hydrolysis, respectively, indicates significant movement of the lower 50-kDa domain. This movement is accompanied by bending at Ile-464 and Gly-466 (chicken S1 sequence numbering), resulting in a rotation (≈20°) and a translation (2.5 Å) of the helix between SH1 and SH2. Although the crystal structure was missing the regulatory domain, extrapolation to the position of the long helix of the heavy chain constituting the regulatory domain suggests that such a movement might alter the position of the regulatory domain relative to the motor domain (5, 31). In a number of studies of isolated myosin heads, significant changes in the hydrodynamic radius were observed (6–8, 32). Such changes have been postulated to generate force (6, 14). Most recently, the displacement of the regulatory domain with respect to the catalytic domain induced by MgADP has been demonstrated in smooth S1 decorating F-actin by three-dimensional electron microscopy reconstruction (9). Similar observations have also been made in skinned fibers with spin-labeled LC2 exchanged into smooth S1 (11). Importantly, domain rearrangement in skeletal muscle was not observed; thus, the reported movement in smooth muscle cannot be the universal conformational change responsible for force generation.

As concluded by both of the above groups, it is not clear whether the observed regulatory movement is responsible for force generation. Our own results do not correlate the structural changes with force generation, because the observed mobility differences between the domains existed in the absence of actin and ATP. However, the presence of a flexible hinge within the myosin head suggests one of the proposed conformational changes during the power stroke must increase the stiffness between the two domains. We might speculate that the hinge stiffness is a function of the attachment to actin filaments and/or the state of ATP hydrolysis products in the active site of myosin as recently implied by Highsmith and Eden (33). The absence of differences between the domains in acto-myosin complexes, implying rigidification of the hinge, hints at this scenario. The possibility of the modulation of this hinge by the intermediate states of the ATPase cycle—in the presence of ADP, aluminofluoride.ATP, and Va.ATP—are being explored. The functional significance of the hinge is not clear, but the observed flexibility might assist in the formation of a stereospecific acto-myosin interface.

Reorientations of the regulatory domain in fibers during contraction has been demonstrated by a number of studies utilizing EPR and time-resolved fluorescence (34–37). These and the finding of direct proportionality between force and length of the regulatory domain (38) suggest that the movement of the regulatory domain is responsible for the power stroke. The capability of relative movement indicates that the regulatory domain can undergo power-stroke transition while the catalytic domain is attached to actin.

In summary, the present ST-EPR study demonstrates that in skeletal myosin filaments the regulatory and catalytic domains of myosin exhibit unequal mobility; the regulatory domain moves slower than the catalytic domain. These results are interpreted as a strong indication that the two domains of the myosin head are connected by a flexible hinge.

Acknowledgments

We thank Drs. Brett Hambly and Ian Trayer for critical reading of the manuscript and many discussions; Dr. Stefan Highsmith for sharing his data prior to publication; Dr. Danuta Szczesna for help with initial experiments; and Liz Fajer and April Adhikari for help in editing the manuscript. This research was sponsored by the National Science Foundation (NSF-IBN-9507477) (to P.F.), the American Heart Association (GIA-9501335) (to P.F.), a student fellowship from the American Heart Association (to B.A.), and the Hungarian National Research Foundation OTKA T-017842 (to K.H).

ABBREVIATIONS

DITC

diisothiocyanate

MSL

N-(1-oxy-2,2,5,5-tetramethyl-4-pyperidinyl)maleimide

IASL

N-(1-oxy-2,2,6,6-tetramethyl-4-piperidinyl)iodoacetamide

InVSL

2-(-oxyl-2,2,5,5-tetramethyl-3-pyrrolin-3-methynyl)indane-1,3-dione

ST-EPR

saturation transfer–EPR

S1

myosin subfragment 1

ELC

essential light chain

Footnotes

§

τR = M(v + h)η/kT, where η denotes the viscosity of the medium; M, the mass of ELC (19, 3 kDa); v, the partial specific volume (0.74 cm3/g); h, the hydration (0.2 g water/g protein); k, the Boltzmann constant; and T, the absolute temperature.

References

  • 1.Cooke R. Curr Opin Cell Biol. 1993;3:590–592. [Google Scholar]
  • 2.Howard J, Spudich J A. Proc Natl Acad Sci USA. 1996;93:4462–4464. [PubMed] [Google Scholar]
  • 3.Huxley A F, Simmons R M. Nature (London) 1971;233:533–538. doi: 10.1038/233533a0. [DOI] [PubMed] [Google Scholar]
  • 4.Rayment I, Holden H M, Whittaker M, Yohn C B, Lorenz M, Holmes K C, Milligan R A. Science. 1993b;261:58–64. doi: 10.1126/science.8316858. [DOI] [PubMed] [Google Scholar]
  • 5.Fisher A J, Smith C A, Thoden J, Smith R, Sutoh K, Holden H M, Rayment I. Biophys J. 1995;68:19s–28s. [PMC free article] [PubMed] [Google Scholar]
  • 6.Highsmith S, Eden D. Biochemistry. 1990;29:4087–4093. doi: 10.1021/bi00469a010. [DOI] [PubMed] [Google Scholar]
  • 7.Wakabayashi K, Tokunaga M, Kohno I, Sugimoto Y, Hamanaka T, Takezawa Y, Wakabayashi T, Amemiya Y. Science. 1992;258:443–447. doi: 10.1126/science.1411537. [DOI] [PubMed] [Google Scholar]
  • 8.Sugimoto Y, Tokunaga M, Takezawa Y, Ikebe M, Wakabayashi K. Biophys J. 1995;68:29s–34s. [PMC free article] [PubMed] [Google Scholar]
  • 9.Jontes J D, Wilson-Kubalek E M, Milligan R A. Nature (London) 1995;378:751–753. doi: 10.1038/378751a0. [DOI] [PubMed] [Google Scholar]
  • 10.Whittaker M, Wilson-Kubalek E M, Smith J E, Faust L, Milligan R A, Sweeney H L. Nature (London) 1995;378:748–751. doi: 10.1038/378748a0. [DOI] [PubMed] [Google Scholar]
  • 11.Gollub J, Cremo C R, Cooke R. Nat Struct Biol. 1996;3:796–802. doi: 10.1038/nsb0996-796. [DOI] [PubMed] [Google Scholar]
  • 12.Rayment I, Rypniewski W R, Schmidt-Base K, Smith R, Tomchick D R, Benning M M, Winkelmann D A, Wesenberg G, Holden H M. Science. 1993;261:50–58. doi: 10.1126/science.8316857. [DOI] [PubMed] [Google Scholar]
  • 13.Highsmith S, Cooke R. Cell Muscle Motil. 1983;4:207–237. [PubMed] [Google Scholar]
  • 14.Cooke R. CRC Crit Rev Biochem. 1986;21:53–118. doi: 10.3109/10409238609113609. [DOI] [PubMed] [Google Scholar]
  • 15.Margossian S, Lowey S. Methods Enzymol. 1982;85:55–71. doi: 10.1016/0076-6879(82)85009-x. [DOI] [PubMed] [Google Scholar]
  • 16.Wagner P D. Methods Enzymol. 1982;85:72–81. doi: 10.1016/0076-6879(82)85010-6. [DOI] [PubMed] [Google Scholar]
  • 17.Pardee J D, Spudich J A. Methods Enzymol. 1982;85:164–181. doi: 10.1016/0076-6879(82)85020-9. [DOI] [PubMed] [Google Scholar]
  • 18.Roopnarine O, Hideg K, Thomas D D. Biophys J. 1993;64:1896–1907. doi: 10.1016/S0006-3495(93)81561-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Adhikari B B, Fajer P G. Biophys J. 1996;70:1872–1880. doi: 10.1016/S0006-3495(96)79752-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Fajer P G, Marsh D. J Magn Reson. 1983;55:205–215. [Google Scholar]
  • 21.Freed J. In: Spin Labeling: Theory and Applications. Berliner L, editor. Vol. 1. New York: Academic; 1976. pp. 53–132. [Google Scholar]
  • 22.Thomas D D, Seidel J C, Gergely J, Hyde J S. J Supramol Struct. 1975;3:376–390. doi: 10.1002/jss.400030410. [DOI] [PubMed] [Google Scholar]
  • 23.Highsmith S, Kretzschmar K M, O’Konski C T, Morales M F. Proc Natl Acad Sci USA. 1977;74:4986–4990. doi: 10.1073/pnas.74.11.4986. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Eads T M, Thomas D D, Austin R H. J Mol Biol. 1984;179:55–81. doi: 10.1016/0022-2836(84)90306-1. [DOI] [PubMed] [Google Scholar]
  • 25.Adhikari B, Somerset J, Hideg K, Fajer P G. Biophys J. 1996;70:A265. (abstr.). [Google Scholar]
  • 26.Robinson B H, Thomann H, Beth A H, Fajer P G, Dalton L R. EPR Studies of Biological Systems. Boca-Raton, FL: CRC; 1985. [Google Scholar]
  • 27.Howard E C, Lindahl K M, Polnaszek C F, Thomas D D. Biophys J. 1993;64:581–593. doi: 10.1016/S0006-3495(93)81417-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28.Broersma S. J Chem Phys. 1960;32:1632–1635. [Google Scholar]
  • 29.Hagerman P J, Zimm B H. Biopolymers. 1981;15:1481–1502. [Google Scholar]
  • 30.Levine R J C, Kensler R W, Yang Z, Stull J T, Sweeney H L. Biophys J. 1996;71:898–907. doi: 10.1016/S0006-3495(96)79293-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Fisher A J, Smith C A, Thoden J B, Smith R, Sutoh K, Holden H M, Rayment I. Biochemistry. 1995;34:8960–8972. doi: 10.1021/bi00028a004. [DOI] [PubMed] [Google Scholar]
  • 32.Suda H, Sugimoto M, Chiba M, Uemura C. Biochem Biophys Res Commun. 1995;211:219–225. doi: 10.1006/bbrc.1995.1799. [DOI] [PubMed] [Google Scholar]
  • 33.Eden D, Highsmith S. Biophys J. 1997;73:952–958. doi: 10.1016/S0006-3495(97)78127-X. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34.Hambly B, Franks K, Cooke R. Biophys J. 1992;63:1306–1313. doi: 10.1016/S0006-3495(92)81717-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35.Irving M, Allen T S, Sabido-David C, Craik J S, Brandmeier B, Kendrick-Jones J, Corrie J E T, Trentham D R, Goldman Y E. Nature (London) 1995;375:688–691. doi: 10.1038/375688a0. [DOI] [PubMed] [Google Scholar]
  • 36.Ling N, Shrimpton C, Sleep J, Kendrick-Jones J, Irving M. Biophys J. 1996;70:1836–1846. doi: 10.1016/S0006-3495(96)79749-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Allen T S, Ling N, Irving M, Goldman Y E. Biophys J. 1996;70:1847–1862. doi: 10.1016/S0006-3495(96)79750-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38.Uyeda T Q P, Abramson P D, Spudich J A. Proc Natl Acad Sci USA. 1996;93:4459–4464. doi: 10.1073/pnas.93.9.4459. [DOI] [PMC free article] [PubMed] [Google Scholar]

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