Comprehensive Physiology Wiley Online Library
For Librarians For Authors Editors About

Kinetics of the Actin-Myosin Interaction

Jeffery W. Walker

10.1002/cphy.cp020106

Source: Supplement 6: Handbook of Physiology, The Cardiovascular System, The Heart

Originally published: 2002

Published online: January 2011

Full Article on Wiley Online Library



Abstract

The sections in this article are:

1 Crossbridges and Sliding Filaments
2 Regulation
3 Myosin and Actomyosin ATPase
3.1 Rates of Specific Steps
3.2 Energetics of Specific Steps
3.3 Cardiac versus Skeletal Actomyosin ATPase
4 The Crossbridge Cycle in Muscle
4.1 Energy Transduction and Muscle Mechanics
4.2 Transient Kinetics in Fibers Using Caged Compounds
4.3 Analysis of Specific Steps
4.4 Cardiac Muscle
5 In Vitro Motility
6 Atomic Structures of Actin and Myosin
6.1 Myosin S1
6.2 Actomyosin
6.3 Comparison of Structural Models to Other Models
7 Recent Progress
8 Regulation
8.1 The Steric Blocking Model
8.2 Kinetic Regulation
8.3 Dual Regulation of the Crossbridge Cycle
8.4 Phosphorylation and Protein Isoform Switching
9 Summary and Concluding Comments
Figure 1.

Muscle fiber ultrastructure. Fibers are comprised of bundles of myofibrils, each containing alternating dark bands (A‐bands) and light bands (I‐bands) when viewed in the light microscope. Illustrated in diagrams below the micrograph is the arrangement of proteins that underlie this banding pattern. The dark band corresponds to the thick myosin filament which has a bare zone or light stripe at its center. The light band corresponds to thin actin filaments which are anchored at the Z‐line in the center of the I‐band. The outer segments of the dark band represent the region of overlap between actin and myosin filaments where crossbridges are located and where productive actin‐myosin interactions occur. A sarcomere is the structure between two Z‐lines, and when the sarcomere shortens the distance between Z‐lines decreases.
Figure 2.

Time course of ATP hydrolysis catalyzed by myosin S1 and actomyosin S1 measured by formation of total Pi. Rapid mixing of ATP with myosin produces a Pi burst in the first few tenths of a second, followed by a slower rate of Pi formation (solid line). The burst is approximately equal in moles to the number of moles of myosin S1 present, and represents the first cycle of ATP hydrolysis bound to myosin S1. Further ATP hydrolysis (and Pi formation) requires product dissociation, which is very slow (t1/2 = 14 s) in the absence of actin. At intermediate actin concentrations, this second phase increases in rate but the overall time course remains distinctly biphasic. At very high actin concentration, the second phase approaches the rate of the initial (burst) phase making the burst less discernible.
Figure 3.

Transduction of chemical free energy into a mechanical process for a simple one step reaction. State A is a non‐force generating state so its basic free energy (GAchem) is independent of displacement. State B is a force generating state whose force is linearly related to displacement (F = κx) and whose mechanical free energy is parabolically related to displacement (GBmech = κx2/2). The dashed line indicates a pathway where all of the free energy is lost as heat and no work is done (as for isolated proteins in solution). The equilibrium constant for the A to B transition, KAB, would be large for the dashed pathway and the reaction would not be readily reversible. The A to B transition can be used to drive a vectorial mechanical process if A is converted to B at some displacement away from the free energy minimum of B (GBchem: where GBmech = 0). The heavy line indicates the pathway for an A to B transition where free energy is efficiently transformed into external work. If after the A to B transition the displacement toward x = 0 were prevented, then the equilibrium constant, KAB, would be small and the reaction would be readily reversible.
Figure 4.

A: A simplified chemical pathway for crossbridges undergoing attachment, force generation, and filament sliding in muscle. Two detached states, M.ATP and M.ADP.Pi, and two attached states, A.M.ADP.Pi and A.M.'ADP, from Scheme 13 are shown. B: Free energy diagram for the two detached x‐independent states, and the two attached x‐dependent states. The heavy line represents the pathway for crossbridges performing work as they move through the cross‐bridge cycle. Vertical steps indicate heat loss; movement along x indicates external work. During isometric contraction, movement along x is prevented so crossbridges become mechanically trapped near position II. These crossbridges would be readily reversible back to A.M.ADP.Pi, M.ADP.Pi and M.ATP (for instance in the presence of high Pi) because of the high free energy content of A.M.'ADP at position II. However, after A.M.'ADP reaches its free energy minimum due to filament sliding or because the proteins are not constrained within the filament (i.e. for ATPase in solution), the A.M.'ADP to A.M.ADP.Pi transition is much less readily reversed. Vectorial force generation and filament sliding require attachment of crossbridges away from their free energy minima. For example, mechanical free energy minima for A.M.ADP.Pi and A.M.'ADP must be displaced relative to one another in the direction appropriate for shortening.
Figure 5.

An idealized force‐velocity relationship (solid line) derived from the normalized Hill equation: V/Vmax = (1‐F/Fo)/(1+(F/Fo)(Fo/α)). This equation describes the observed inverse relationship between force (or load) and velocity with a characteristic curvature (Fo/α) for each fiber type. The left extreme of the curve is dominated by the action of positively strained crossbridges, the right side by detachment of negatively strained crossbridges, and the middle by dynamic mixtures of positively and negatively strained cycling bridges. A power‐load relationship (dashed line) derived from the idealized forced‐velocity data shows maximum power at 0.3Vmax. The power‐load curve and efficiency also depend upon the Fo/α factor for a given fiber type 49,112.
Figure 6.

Schematic representation of crossbridges in three states. I. Detached or weakly attached. In this state the crossbridge can assume many distinct orientations due to a flexible hinge where an elastic spring element attaches to the globular head. II. Strongly attached positively strained. This state would occur during isometric contraction after attachment, Pi release, and rotation of the crossbridge to stretch the spring. III. If the filaments are released the spring will recoil causing filament displacement. This state would occur if filaments were to slide beyond the mechanical equilibrium position to a position where the spring is compressed. This is a negatively strained crossbridge that impedes further sliding. Approximate locations of I-III on the free energy diagram are given in Figure 4B.
Figure 7.

Structure and photolysis half‐times for three caged compounds commonly used in skinned fiber studies. Approximate half‐times are listed for the rate‐limiting photochemical reactions under near physiological conditions, pH 7, 21°C. The version of caged Ca2+ illustrated is nitrophenyl EGTA 20.
Figure 8.

Close coupling between force generation and Pi release in skinned cardiac myocytes. A: Isometric force development after photochemical elevation of free Ca2+ within the filament lattice. The time course prior to “slack” shows that both the rate and extent of force development were altered by the presence of 10 mM added Pi (b) compared to no added Pi (a). Pi depressed the amplitude of the force response and accelerated the rate of approach to the final amplitude, consistent with Scheme 14. B: A complimentary experiment in which Pi was produced rapidly within isometric Ca2+‐activated myocytes. Increasing concentrations of photoreleased Pi, (a) 0.5 mM Pi, (b) 1 mM Pi and (c) 2.7 mM Pi, caused both a greater and more rapid depression of isometric force, consistent with Scheme 14. For comparison, Pi at 0.5 to 10 mM has no effect on the ATPase of isolated actomyosin. Data from 2 with permission.
Figure 9.

Elementary transitions in single myosin molecules. A: Experimental arrangement for optical trap measurements of unitary force and displacement by actomyosin. The actin filament is attached at each end to a bead (dark spheres). Each bead is controlled by a laser trap. The actin filament is lowered onto another bead (white sphere) coated with a low density of myosin molecules. From 23 with permission. B, C: Records of force and displacement produced by individual V1 and V1 cardiac myosin isoforms. The unitary force and displacements are similar for V1 and V3, but V3 events are prolonged. Verticle scale bars are 2 pN and 20 nm, respectively. D: Schematic of unitary events in cardiac myosin. In principle, myosins can differ by amplitude of unitary force or displacement on the y‐axis, by mean duration of attached strong binding states, or by duration of detached or weak binding states. Dotted lines illustrate time‐averaged force or displacement.

From 91 with permission
Figure 10.

Domains and clefts of myosin S1. A: Outlines of major domains are superimposed on a ribbon diagram taken from the crystal structure of chicken myosin S1 76,78. Also shown are the nucleotide cleft (vertical arrow) and actin site cleft (horizontal arrow). B: Schematic representation of myosin S1 including 20K (long helix), 25K, 50K, LC1, LC2, and an elastic element. In the nucleotide‐free state (left), the nucleotide site at the 25K-50K interface is open, but the actin interaction site at the 20K-50K interface is closed and S1 is tightly bound to actin. ATP binding in the nucleotide pocket results in large changes in the actin interaction site causing a greatly weakened interaction between myosin S1 and actin (right). Dissociation of Pi out of the actin site cleft permits the 20K-50K interface to close and strong bonds between S1 and actin to reform (bottom). An appropriate change in the angle of the 10 nm extended arm of the 20K domain, stabilized by light chains, could stretch an elastic element or displace the actin and myosin filaments relative to one another.


Figure 1.

Muscle fiber ultrastructure. Fibers are comprised of bundles of myofibrils, each containing alternating dark bands (A‐bands) and light bands (I‐bands) when viewed in the light microscope. Illustrated in diagrams below the micrograph is the arrangement of proteins that underlie this banding pattern. The dark band corresponds to the thick myosin filament which has a bare zone or light stripe at its center. The light band corresponds to thin actin filaments which are anchored at the Z‐line in the center of the I‐band. The outer segments of the dark band represent the region of overlap between actin and myosin filaments where crossbridges are located and where productive actin‐myosin interactions occur. A sarcomere is the structure between two Z‐lines, and when the sarcomere shortens the distance between Z‐lines decreases.



Figure 2.

Time course of ATP hydrolysis catalyzed by myosin S1 and actomyosin S1 measured by formation of total Pi. Rapid mixing of ATP with myosin produces a Pi burst in the first few tenths of a second, followed by a slower rate of Pi formation (solid line). The burst is approximately equal in moles to the number of moles of myosin S1 present, and represents the first cycle of ATP hydrolysis bound to myosin S1. Further ATP hydrolysis (and Pi formation) requires product dissociation, which is very slow (t1/2 = 14 s) in the absence of actin. At intermediate actin concentrations, this second phase increases in rate but the overall time course remains distinctly biphasic. At very high actin concentration, the second phase approaches the rate of the initial (burst) phase making the burst less discernible.



Figure 3.

Transduction of chemical free energy into a mechanical process for a simple one step reaction. State A is a non‐force generating state so its basic free energy (GAchem) is independent of displacement. State B is a force generating state whose force is linearly related to displacement (F = κx) and whose mechanical free energy is parabolically related to displacement (GBmech = κx2/2). The dashed line indicates a pathway where all of the free energy is lost as heat and no work is done (as for isolated proteins in solution). The equilibrium constant for the A to B transition, KAB, would be large for the dashed pathway and the reaction would not be readily reversible. The A to B transition can be used to drive a vectorial mechanical process if A is converted to B at some displacement away from the free energy minimum of B (GBchem: where GBmech = 0). The heavy line indicates the pathway for an A to B transition where free energy is efficiently transformed into external work. If after the A to B transition the displacement toward x = 0 were prevented, then the equilibrium constant, KAB, would be small and the reaction would be readily reversible.



Figure 4.

A: A simplified chemical pathway for crossbridges undergoing attachment, force generation, and filament sliding in muscle. Two detached states, M.ATP and M.ADP.Pi, and two attached states, A.M.ADP.Pi and A.M.'ADP, from Scheme 13 are shown. B: Free energy diagram for the two detached x‐independent states, and the two attached x‐dependent states. The heavy line represents the pathway for crossbridges performing work as they move through the cross‐bridge cycle. Vertical steps indicate heat loss; movement along x indicates external work. During isometric contraction, movement along x is prevented so crossbridges become mechanically trapped near position II. These crossbridges would be readily reversible back to A.M.ADP.Pi, M.ADP.Pi and M.ATP (for instance in the presence of high Pi) because of the high free energy content of A.M.'ADP at position II. However, after A.M.'ADP reaches its free energy minimum due to filament sliding or because the proteins are not constrained within the filament (i.e. for ATPase in solution), the A.M.'ADP to A.M.ADP.Pi transition is much less readily reversed. Vectorial force generation and filament sliding require attachment of crossbridges away from their free energy minima. For example, mechanical free energy minima for A.M.ADP.Pi and A.M.'ADP must be displaced relative to one another in the direction appropriate for shortening.



Figure 5.

An idealized force‐velocity relationship (solid line) derived from the normalized Hill equation: V/Vmax = (1‐F/Fo)/(1+(F/Fo)(Fo/α)). This equation describes the observed inverse relationship between force (or load) and velocity with a characteristic curvature (Fo/α) for each fiber type. The left extreme of the curve is dominated by the action of positively strained crossbridges, the right side by detachment of negatively strained crossbridges, and the middle by dynamic mixtures of positively and negatively strained cycling bridges. A power‐load relationship (dashed line) derived from the idealized forced‐velocity data shows maximum power at 0.3Vmax. The power‐load curve and efficiency also depend upon the Fo/α factor for a given fiber type 49,112.



Figure 6.

Schematic representation of crossbridges in three states. I. Detached or weakly attached. In this state the crossbridge can assume many distinct orientations due to a flexible hinge where an elastic spring element attaches to the globular head. II. Strongly attached positively strained. This state would occur during isometric contraction after attachment, Pi release, and rotation of the crossbridge to stretch the spring. III. If the filaments are released the spring will recoil causing filament displacement. This state would occur if filaments were to slide beyond the mechanical equilibrium position to a position where the spring is compressed. This is a negatively strained crossbridge that impedes further sliding. Approximate locations of I-III on the free energy diagram are given in Figure 4B.



Figure 7.

Structure and photolysis half‐times for three caged compounds commonly used in skinned fiber studies. Approximate half‐times are listed for the rate‐limiting photochemical reactions under near physiological conditions, pH 7, 21°C. The version of caged Ca2+ illustrated is nitrophenyl EGTA 20.



Figure 8.

Close coupling between force generation and Pi release in skinned cardiac myocytes. A: Isometric force development after photochemical elevation of free Ca2+ within the filament lattice. The time course prior to “slack” shows that both the rate and extent of force development were altered by the presence of 10 mM added Pi (b) compared to no added Pi (a). Pi depressed the amplitude of the force response and accelerated the rate of approach to the final amplitude, consistent with Scheme 14. B: A complimentary experiment in which Pi was produced rapidly within isometric Ca2+‐activated myocytes. Increasing concentrations of photoreleased Pi, (a) 0.5 mM Pi, (b) 1 mM Pi and (c) 2.7 mM Pi, caused both a greater and more rapid depression of isometric force, consistent with Scheme 14. For comparison, Pi at 0.5 to 10 mM has no effect on the ATPase of isolated actomyosin. Data from 2 with permission.



Figure 9.

Elementary transitions in single myosin molecules. A: Experimental arrangement for optical trap measurements of unitary force and displacement by actomyosin. The actin filament is attached at each end to a bead (dark spheres). Each bead is controlled by a laser trap. The actin filament is lowered onto another bead (white sphere) coated with a low density of myosin molecules. From 23 with permission. B, C: Records of force and displacement produced by individual V1 and V1 cardiac myosin isoforms. The unitary force and displacements are similar for V1 and V3, but V3 events are prolonged. Verticle scale bars are 2 pN and 20 nm, respectively. D: Schematic of unitary events in cardiac myosin. In principle, myosins can differ by amplitude of unitary force or displacement on the y‐axis, by mean duration of attached strong binding states, or by duration of detached or weak binding states. Dotted lines illustrate time‐averaged force or displacement.

From 91 with permission


Figure 10.

Domains and clefts of myosin S1. A: Outlines of major domains are superimposed on a ribbon diagram taken from the crystal structure of chicken myosin S1 76,78. Also shown are the nucleotide cleft (vertical arrow) and actin site cleft (horizontal arrow). B: Schematic representation of myosin S1 including 20K (long helix), 25K, 50K, LC1, LC2, and an elastic element. In the nucleotide‐free state (left), the nucleotide site at the 25K-50K interface is open, but the actin interaction site at the 20K-50K interface is closed and S1 is tightly bound to actin. ATP binding in the nucleotide pocket results in large changes in the actin interaction site causing a greatly weakened interaction between myosin S1 and actin (right). Dissociation of Pi out of the actin site cleft permits the 20K-50K interface to close and strong bonds between S1 and actin to reform (bottom). An appropriate change in the angle of the 10 nm extended arm of the 20K domain, stabilized by light chains, could stretch an elastic element or displace the actin and myosin filaments relative to one another.

References
 1. Araujo, A. and J. W. Walker. Kinetics of tension development in skinned cardiac myocytes measured by photorelease of Ca2+ Am. J. Physiol. 267 Heart Circ. Physiol. 36: H1643-H1653, 1994.
 2. Araujo, A. and J. W. Walker. Phosphate release and force generation in cardiac myocytes investigated with caged phosphate and caged Ca2+. Biophys. J. 70: 2316-2326, 1996.
 3. Baker, J. E., I. Brust‐Mascher, S. Ramachandran, L. E. LaConte, and D. D. Thomas. A large and distinct rotation of the myosin light chain domain occurs upon muslce contraction. Proc. Nat. Acad. Sci. U.S.A. 95: 2720-2722, 1998.
 4. Bagshaw, C. R. Muscle Contraction, London: Chapman Hall, 1993.
 5. Barany, M. ATPase activity of myosin correlated with speed of muscle shortening. J. Gen. Physiol. 50: 197-218, 1967.
 6. Barsotti, R. J. and M. A. Ferenczi. Kinetics of ATP hydrolysis and tension production in skinned cardiac muscle of the guinea pig. J. Biol. Chem. 263: 16750-16756, 1988.
 7. Bowater, R. and J. Sleep. Demembranated muscle fibers catalyze a more rapid exchange between phosphate and adenosine triphosphate than actomyosin subfragment 1. Biochemistry 27: 5314-5323, 1988.
 8. Brenner, B. Effect of Ca2+ on cross‐bridge turnover kinetics in skinned single rabbit psoas fibers: Implications for regulation of muscle contraction. Proc. Natl. Aced. Sci. U.S.A. 85: 3265-3269, 1988.
 9. Burton, K. Myosin step size: estimates from motility assays and shortening muscle. J. Muscle Res. Cell Motil. 13: 590-607, 1992.
 10. Chalovich, J. M. and E. Eisenberg. Inhibition of actomyosin ATPase activity by troponin‐tropomyosin without blocking the binding of myosin to actin. J. Biol. Chem. 257: 2432-2437, 1982.
 11. Cooke, R. Actomyosin interaction in striated muscle. Physiol. Rev. 77: 671-697, 1997.
 12. Dantzig, J. A. and Y. E. Goldman. Suppression of muscle contraction by vanadate. J. Gen. Physiol. 86: 305-327, 1985.
 13. Dantzig, J. A., M. A. Hibberd, D. R. Trentham, and Y. E. Goldman. Crossbridge kinetics in the presence of MgADP investigated by photolysis of caged ATP in rabbit psoas muscle fibers. J. Physiol. (Lond.) 432: 639-680, 1991.
 14. Dantzig, J. A., Y. E. Goldman, N. C. Millar, J. Laktis, and E. Homsher. Reversal of the cross‐bridge force‐generating transition by photogeneration of phosphate in rabbit psoas muscle fibers. J. Physiol. (Lond.) 451: 247-278, 1992.
 15. Dantzig, J. A., J. W. Walker, D. R. Trentham, and Y. E. Goldman. Relaxation of muscle fibers with ATP(γS) and by laser photolysis of caged ATP(γS): Evidence for Ca2+ dependent affinity of rapidly detaching zero force cross‐bridges. Proc. Natl. Acad. Sci. U.S.A. 85: 6716-6720, 1988.
 16. Dominquez, R., Y. Freyzon, K. M. Trybus, and C. Cohen. Crystal structure of a vertebrate smooth muscle myosin motor domain and its complex with the essential light chain: visualization of the pre‐power stroke state. Cell 94: 559-571, 1998.
 17. Ebashi, S. Calcium ions and muscle contraction. Nature 240: 217-218, 1972.
 18. Eisenberg, E. and T. L. Hill. Muscle contraction and free energy transduction in biological systems. Science 227: 999-1006, 1985.
 19. Ellis‐Davies, G. C. R. and J. A. Kaplan. Nitrophenyl EGTA, a photolabile chelator that selectively binds Ca2+ with high affinity and rapidly releases it upon photolysis. Proc. Natl. Acad. Sci. U.S.A. 91: 187-191, 1994.
 20. Fenn, W. O. A quantitative comparison between the energy liberated and the work performed by the isolated sartorius of the frog. J. Physiol. (Lond.) 58: 175-203, 1923.
 21. Ferenczi, M. A., E. Homsher, and D. R. Trentham. The kinetics of magnesium adenosine triphosphate cleavage in skinned muscle fibres of the rabbit. J. Physiol. (Lond.) 352: 575-599, 1984.
 22. Ferenczi, M. A. Phosphate burst in permeable muscle fibers of the rabbit. Biophys. J. 50: 471-477, 1986.
 23. Finer, J. T., R. A. Simmons, and J. A. Spudich. Single myosin molecule mechanics: piconewton forces and nanometre steps. Nature 368: 113-119, 1994.
 24. Fisher, A. J., C. A. Smith, J. Thoden, R. Smith, K. Sutoh, H. Holden, and I. Rayment. Structural studies of myosin:nucleotide complexes: a revised model for the molecular basis of muscle contraction. Biophys. J. 68: 19s-26s, 1995.
 25. Ford, L. E., A. F. Huxley, and R. M. Simmons. Tension responses to sudden length changes in stimulate from muscle fibers near slack length. J. Physiol. (Lond.) 269: 441-515, 1977.
 26. Ishijima, A., H. Kojima, T. Funatsu, K. Tokunaga, H. Higuchi, H. Tanaka, and T. Yanagida. Simultaneous observation of individaul ATPase and mechanical events by a singly myosin molecule during interation with actin. Cell 92: 161-171, 1998.
 27. Goodson, H. V. and J. A. Spudich. Molecular evolution of the myosin family: relationships derived from comparisons of amino acid sequences. Proc. Natl. Acad. Sci. USA 90: 659-663, 1993.
 28. Goldman, Y. E., M. G. Hibberd, and D. R. Trentham. Relaxation of rabbit psoas muscle fibers from rigor by photochemical generation of adenosine‐5'‐triphosphate. J. Physiol. 354: 577-604, 1984.
 29. Goldman, Y. E. Kinetics of the actomyosin ATPase in muscle fibers. Annu. Rev. Physiol. 49: 637-654, 1987.
 30. Goldman, Y. E. Wag the tail: structural dynamics of actomyosin. Cell 93: 1-4, 1998.
 31. Gordon, A. M., A. F. Huxley, and F. J. Julian. The variation in isometric tension with sarcomere length in vertebrate muscle fibers. J. Physiol. 184: 170-192, 1966.
 32. Guilford, W. H., D. E. Dupuis, G. Kennedy, J. Wu, J. B. Patlak, and D. M. Warshaw. Smooth muscle and skeletal muscle myosins produce similar unitary forces and displacements in the laser trap. Biophys. J. 72: 1006-1021, 1997.
 33. Gulick, A. M. and I. Rayment. Structural studies of myosin II: communication between distant protein domains. Bioessays 19: 561-569.
 34. Hibberd, M. G. and D. R. Trentham. Relationships between chemical and mechanical events during muscular contraction. Annu. Rev. Biophys. Biophys. Chem. 15: 119-161, 1986.
 35. Highsmith, S. Lever arm model of force generation by actinmyosin‐ATP. Biochemistry 38: 9791-9797, 1999.
 36. Holmes, K. C., D. Popp, W. Gebhard, and W. Kabsch. Atomic model of the actin filament. Nature 347: 44-49, 1990.
 37. Holmes, K. C. The swinging lever‐arm hypothesis of muscle contraction. Curr. Biol. 7: R112-R118, 1997.
 38. Homsher, E. and N. C. Millar. Caged compounds and striated muscle contraction. Annu. Rev. Physiol. 52: 875-896, 1990.
 39. Houdusse, A., V. N. Kalbokis, D. Himmel, A. G. Szent‐Gyorgyi, and C. Cohen. Atomic structure of scallop myosin subfragment S1 complexed with MgADP: a novel comformation of the myosin head. Cell 97: 459-470, 1999.
 40. Huxley, A. F. Muscle structure and theories of contraction. Prog. Biophys. Biophys. Chem. 7: 255-318, 1957.
 41. Huxley, A. F. and R. Niedergerke. Structural changes in muscle during contraction. Nature 173: 971-973, 1954.
 42. Huxley, A. F. and R. M. Simmons. Proposed mechanism of force generation in muscle fibers. Nature 233: 533-538, 1971.
 43. Huxley, A. F. Reflections on Muscle. Princeton, NJ: Princeton University Press, 1980.
 44. Huxley, H. E. and J. Hanson. Changes in the cross‐striations of muscle during contraction and stretch and their structural interpretation. Nature 173: 973-976, 1954.
 45. Huxley, H. E. The mechanism of muscular contraction. Science 164: 1356-1366, 1969.
 46. Huxley, H. E. Structural changes in actin‐ and myosin‐containing filaments during contraction. Cold Spring Harbor Symp. Quant. Biol. 37: 361-376, 1973.
 47. Irving, M., V. Lombardi, G. Piazzesi, and M. A. Ferenczi. Myosin head movements are synchronous with the elementary force‐generating process in muscle. Nature 357: 156-158, 1992.
 48. Irving, M., T. S. C. Allen, C. Sabido‐David, J. S. Craik, B. Brandmeier, J. Kendrick‐Jones, J. E. T. Corrie, D. R. Trentham, and Y. E. Goldman. Tilting of the light‐chain region of myosin during step length changes and active force generation in skeletal muscle. Nature 375: 688-691, 1995.
 49. Josephson, R. K. Contraction dynamics and power output of skeletal muscle. Annu. Rev. Physiol. 55: 527-54, 1993.
 50. Julian, F. J. and M. R. Sollins. Variation of muscle stiffness with force at increasing speeds of shortening. J. Gen. Physiol. 66: 287-302. 1975.
 51. Kabsch, W., H. G. Mannherz, D. Suck, E. F. Pai, and K. C. Holmes. Atomic structure of the actin: DNasel complex. Nature 347: 21-22, 1990.
 52. Kawai, M., Y. Saeki, and Y. Zhao. Cross‐bridge scheme and the kinetic constants of elementary steps deduced from chemically skinned papillary and trabecular muscles of the ferret. Circ. Res. 73: 35-50, 1993.
 53. Kitamura, K., M. Tokunaga, A. H. Iwane, and T. Yanagida. A single myosin head moves along actin filaments with regular steps of 5.3 nanometers. Nature 397: 129-134.
 54. Kress, M., H. E. Huxley, A. R. Farqui, and J. Hendrix. Structural changes during activation of frog muscle studied by time‐resolved x‐ray diffraction. J. Mol. Biol. 188: 325-342, 1985.
 55. Kushmerick, M. J. and R. E. Davies. The chemical energetics of muscle contraction. II. The chemistry, efficiency and power of maximally working sartorius muscle. Proc. R. Soc. London B 174: 315-353, 1969.
 56. Lankford, E. B., N. D. Epstein, L. Fananpazir, and H. L. Sweeney. Abnormal contractile properties of muscle fibers expressing beta‐myosin heavy chain gene mutations in patients with hypertrophic cardiomyopathy. J. Clin. Invest. 95: 1409-1414, 1995.
 57. Lauzon, A. M., M. J. Tyska, A. S. Rovner, Y. Freyon, D. M. Warshaw, and K. M. Trybus. A 7‐amino acid insert in the heavy chain nucleotide binding loop alters the kinetics of smooth muscle myosin in the laser trap. J. Muscle Cell Res. Cell Motil. 19: 825-837, 1998.
 58. Lehman, W., P. Vibert, P. Uman, and R. Craig. Steric blocking by tropomyosin visualized in relaxed vertebrate muscle filaments. J. Mol. Biol. 251: 191-196, 1995.
 59. Lionne, C., M. Brune, M. R. Webb, F. Travers, and T. Barman. Time resolved measurements show that phosphate release is the rate limiting step on myofibrillar ATPase. FEBS Lett. 364: 59-62, 1995.
 60. Lowey, S., G. S. Waller, and K. M. Trybus. Skeletal muscle light chains are essential for physiological speeds of shortening. Nature 365: 454-456, 1993.
 61. Lu, Z., R. L. Moss, and J. W. Walker. Tension transients initiated by photogeneration of MgADP in skinned skeletal muscle fibers. J. Gen. Physiol. 101: 867-888, 1993.
 62. Lymn, R. W. and E. Taylor. Mechanism of adenosine triphosphate hydrolysis by actomyosin. Biochemistry 10: 4617-4624, 1971.
 63. Ma, Y. Z. and E. W. Taylor. Kinetic mechanism of myofibril ATPase. Biophys. J. 66: 1542-1553, 1994.
 64. McKillop, D. F and M. A. Geeves. Regulation of the actomyosin subfragment 1 interaction by troponin/tropomyosin. Biophys. J. 65: 693-701, 1993.
 65. Martin, H. and R. J. Barsotti. Relaxation from rigor of skinned trabeculae of the guinea pig induced by laser photolysis of caged ATP. Biophys. J. 66: 1115-1128, 1994.
 66. Martin, H. and R. J. Barsotti. Activation of skinned trabeculae of the guinea pig induced by laser photolysis of caged ATP. Biophys. J. 67: 1933-1941, 1994.
 67. Metzger, J. M., M. L. Greaser, and R. L. Moss. Variations in cross‐bridge attachment rate and tension with phosphorylation of myosin in skinned skeletal muscle fibers. J. Gen. Physiol. 93: 855-883, 1989.
 68. Millar, N. C. and E. Homsher. The effect of phosphate and calcium on force generation in glycerinated rabbit skeletal muscle fibers. J. Biol. Chem. 265: 20234-20240, 1990.
 69. Milligan, R. A. Protein‐protein interactions in the rigor actomyosin complex. Proc. Natl. Acad. Sci. U.S.A. 93: 21-26, 1996.
 70. Milligan, R. A., M. Wittaker, and D. Safer. Molecular structure of F‐actin and location of surface binding sites. Nature 348: 217-221, 1990.
 71. Molloy, J. E., J. E. Burns, J. Kendrick‐Jones, R. T. Tregear and D. C. S. White. Force and movement produced by a single myosin head. Nature 378: 209-212, 1995.
 72. Murphy, C. T. and J. A. Spudich. The sequence of the myosin 50-20K loop affects myosins affinity for actin throughout the actin‐myosin ATPase cycle and its maximum ATPAse activity. Biochemistry 38: 3785-3792, 1999.
 73. Parry, D. A. D. and J. M. Squire. Structural role of tropomyosin in muscle regulation: analysis of the X‐ray diffraction patterns from relaxed and contracting muscle. J. Mol. Biol. 75: 33-55, 1973.
 74. Pope, B., J. F. Y. Hoh, and A. Weeds. The ATPase activities of rat cardiac myosin isoenzymes. FEBS Lett. 118: 205-208, 1980.
 75. Rall, J. A. Energetic aspects of skeletal muscle contraction: implications of fiber types. Exerc. Sports Sci. Rev. 13: 33-74, 1985.
 76. Rayment, I., W. R. Rypniewski, K. Schmidt‐Base, R. Smith, D. R. Tomchick, M. M. Benning, D. A. Winkelman, G. Wesenberg, and H. M. Holden. Three dimensional structure of myosin subfragment 1: a molecular motor. Science 261: 35-36, 1993.
 77. Rayment, I., H. M. Holden, M. Wittaker, C. B. Yohn, M. Lorenz, K. C. Holmes, and R. A. Milligan. Structure of the actinmyosin complex and its implications for muscle contraction. Science 261: 58-65, 1993.
 78. Rayment, I. and H. Holden. The three dimensional structure of a molecular motor. Trends Biochem. Sci. 19: 129-134, 1994.
 79. Rayment, I., H. M. Holden, J. R. Sellers, L. Fananapazir, F. Epstein. Structural interpretation of the mutations in the β‐cardiac myosin that have been implicated in familial hypertropic cardiomyopathy. Proc. Natl. Acad. Sci. U.S.A. 92: 3864-3868, 1995.
 80. Reedy, M., K. C. Holmes, and R. T. Tragear. Induced changes in orientation of the cross‐bridges of glycerinated insect flight muscle. Nature 207: 1276-1280, 1965.
 81. Rome, L. C., C. Cook, D. A. Syme, M. A. Connaughton, M. Ashley‐Ross, A. Klimov, B. Tikunov and Y. E. Goldman. Trading force for speed: why superfast crossbridge kinetics leads to superlow forces. Proc. Nat. Acad. Sci. U.S.A. 96: 5826-5831, 1999.
 82. Rosenfeld, S. S. and E. W. Taylor. The ATPase mechanism of skeletal and smooth muscle acto‐subfragment 1. J. Biol. Chem. 259: 11908-11918, 1984.
 83. Rosenfeld, S. S. and E. W. Taylor. The mechanism of regulation of acto‐subfragment 1 ATPase. J. Biol. Chem. 262: 9984-9993, 1987.
 84. Ruppel, K. M., M. Lorenz, and J. A. Spudich. Myosin structure/function: a combined mutagenesis‐crystallographic approach. Curr. Opin. in Struct. Biol. 5: 181-186, 1995.
 85. Schroder, R. R., D. J. Manstein, W. Jahn, H. Holden, I. Rayment, K. C. Holmes, and J. A. Spudich. Three‐dimensional atomic model of F‐actin decorated with Dictyostelium myosin S1. Nature 364: 171-174, 1993.
 86. Siemankowski, R. F. and H. D. White. Kinetics of the interaction between actin, ADP and cardiac myosin S1. J. Biol. Chem. 259: 5045-5053. 1984.
 87. Siemankowski, R. F., M. O. Wiseman, and H. D. White. ADP dissociation from actomyosin subfragment 1 is sufficiently slow to limit the unloaded shortening velocity in muscle. Proc. Natl. Acad. Sci. U.S.A. 82: 658-666, 1985.
 88. Simmons, R. Molecular motors: single‐molecule mechanics. Curr. Biology 6: 392-394.
 89. Sleep, J. A. and R. L. Hutton. Exchange between inorganic phosphate and adenosine 5'‐triphosphate in the medium by actomyosin subfragment 1. Biochemistry 19: 1276-1283, 1980.
 90. Solaro, R. J. and H. M. Rarick. Troponin and tropomyosin: proteins that switch on and tune in the activity of cardiac myofilaments. Circ. Res. 83: 471-480, 1998.
 91. Spudich, J. A. How molecular motors work. Nature 372: 515-518, 1994.
 92. Suguira, S. N. Kobayakawa, H. Fujita, H. Yamashita, S. Monomura, S. Chaen, M. Omata, and H. Sugi. Comparison of unitary displacements and forces between 2 cardiac myosin isoforms by the optical trap technique: molecular basis for cardiac adaptation. Circ. Res. 82: 1029-1034, 1998.
 93. Sweeney, H. L. and E. L. F. Holzbar. Mutational analysis of motor proteins. Annu. Rev. Physiol. 58: 751-792, 1996.
 94. Swartz, D. R. and R. L. Moss. Influence of a strong binding myosin analogue on calcium sensitive mechanical properties of skinned skeletal muscle fibers. J. Biol. Chem. 267: 20497-20506, 1992.
 95. Taylor, E. W. Mechanism of actomyosin ATPase and the problem of muscle contraction. CRC Crit. Rev. Biochem. 6: 103-164, 1979.
 96. Thirlwell, H., J. E. T. Corrie, G. P. Reid, D. R. Trentham, and M. A. Ferenczi. Kinetics of relaxation from rigor of permeabilized fast‐twitch skeletal fibers from the rabbit using a novel caged ATP and apyrase. Biophys. J. 67: 2346-2447, 1994.
 97. Thomas, D. D., S. Ramachandran, O. Roopnarine, D. W. Hayden, and E. Ostap. The mechanism of force generation in muscle: a disorder‐to‐order transition coupled to internal structural change. Biophys. J. 68: 135s-141s, 1995.
 98. Toyoshima, Y. Y., S. J. Kron, E. M. McNally, K. R. Niebling, C. Toyoshima, and J. A. Spudich. Myosin subfragment 1 is sufficient to move actin filaments in vitro. Nature 328: 536-539, 1987.
 99. Trentham, D. R., J. F. Eccleston and C. R. Bagshaw. Kinetic analysis of ATPase mechanisms. Q. Rev. Biophys. 9: 217-281, 1976.
 100. Tyska, M. J., D. E. Dupuis, W. H. Guilford, J. B. Patlak, G. S. Waller, K. M. Trybus, D. M. Warshaw, and S. Lowey. Two heads of myosin are better than one for generating force and motion. Proc. Nat. Acad. Sci. U.S.A. 96: 4402-4407, 1999.
 101. Uyeda, T. Q. and J. A. Spudich. A functional recombinant myosin II lacking a regulatory light chain binding site. Science 262: 1867-1870, 1993.
 102. Uyeda, T. Q., K. M. Ruppel, and J. A. Spudich. Enzymatic activities correlate with chimeric substitutions at the actin‐binding face of myosin. Nature 368: 567-569. 1994.
 103. VanBuren, P., G. S. Waller, D. E. Harris, K. M. Trybus, D. M. Warshaw, and S. Lowey. The essential light chain is required for full force production by skeletal muscle myosin. Proc. Natl. Acad. Sci. U.S.A. 91: 12403-12407, 1994.
 104. VanBuren, P., D. E. Harris, N. R. Alpert and D. M. Warshaw. Cardiac V1 and V3 myosins differ in their hydrolytic and mechanical activities in vitro. Circ. Res. 77: 439-444, 1995.
 105. Veigel, C., L. M. Coluccio, J. D. Contes, J. C. Sparrow, R. A. Milligan, and J. E. Molloy. The motor protein myosin I produces its working stroke in two steps. Nature 398: 530-533, 1999.
 106. Walker, J. W., Z. Lu and R. L. Moss. Effects of Ca2+ on the kinetics of phosphate release in skeletal muscle. J. Biol. Chem. 267: 2459-2466, 1992.
 107. Walker, J. W., G. Reid, J. A. McCray, and D. R. Trentham. Photolabile 1‐(2‐nitrophenyl)ethyl phosphate esters of adenine nucleotide analogues. Synthesis and mechanism of photolysis. J. Am. Chem. Soc. 110: 7170-7177, 1988.
 108. Webb, M. R., M. G. Hibberd, Y. E. Goldman, and D. R. Trentham. Oxygen exchange between Pi in the medium and water during ATP hydrolysis mediated by skinned fibers from rabbit skeletal muscle. J. Biol. Chem. 261: 15557-15564, 1986.
 109. Wells, A. L., A. W. Lin, L. Q. Chen, D. Safer, S. M. Cain, T. Hasson, B. O. Caragher, R. A. Milligan and H. L. Sweeney. Myosin VI is an actin‐based motor that moves backwards. Nature 401: 431-432, 1999.
 110. White, H. D. and E. W. Taylor. Energetics and mechanism of actomyosin adenosine triphosphatase. Biochemistry 15: 5818-5826, 1976.
 111. Wittaker, M., E. M. Wilson‐Kubalik, J. E. Smith, L. Faust, R. A. Milligan and H. L. Sweeney. A 35‐A movement of smooth muscle myosin on ADP release. Nature 378: 748-757, 1995.
 112. Woledge, R. C., N. A. Curtin, and E. Homsher. Energetic Aspects of Muscle Contraction London: Academic Press, 1985.
 113. Yount, R. G., D. Lawson, and I. Rayment. Is myosin a “back door” enzyme? Biophys. J. 68: 44s-47s, 1995.
 114. Zhao, Y. and M. Kawai. Kinetic and thermodynamic studies of the crossbridge cycle in rabbit psoas muscle fibers. Biophys. J. 67: 1655-1658, 1994.

Contact Editor

Submit a note to the editor about this article by filling in the form below.

* Required Field

Article Title: Kinetics of the Actin-Myosin Interaction
 

How to Cite

Jeffery W. Walker. Kinetics of the Actin-Myosin Interaction. Compr Physiol 2011, Supplement 6: Handbook of Physiology, The Cardiovascular System, The Heart: 240-263. First published in print 2002. doi: 10.1002/cphy.cp020106