Actin-Myosin Interaction

Structural model: Both F-actin and myosin are large protein molecules whose interaction takes place at multiple sites in a very specific manner. From the independent X-ray structure of G-actin (Fig. A5) and S1 (Fig. M9) and from the electron density maps of actin filaments decorated with S1, the three-dimensional structure of actoS1 was reconstructed as shown on Fig. AM1.

Fig. AM1. Three-dimensional atomic model of F-actin combined with myosin subfragment 1. (From , Geeves and Holmes, with permission from the Annual Review of Biochemistry, vol. 68,1999, by Annual Reviews, http://www.AnnualReviews.org). Ribbon representation of the myosin-actin interaction. The ribbon on the left corresponds to myosin subfragment 1; an F-actin double helix consisting of 5 actin globules is shown on the right.

Contact sites: As discussed in connection with Fig. A5, the G-actin molecule has four subdomains, most of the amino acids that are involved in the interaction with S1 are located in the subdomain 1.

As discussed in connection with the three-dimensional structure of S1, the first helix of the 20-kDa fragment and the lower and upper domains of the 50-kDa fragment contain the actin-binding site of S1.

The first contact involves positively charged amino acid residues of S1 at the 20-50-kDa junction and negatively charged amino acid residues of actin subdomain 1. This is a non-specific electrostatic contact; it constitutes the weak binding between S1 and actin.

The second contact involves hydrophobic residues of both S1 and actin. This is a stereospecific interaction. The lower domain of the 50-kDa fragment of S1 is linked to residues mainly from subdomain 1 of actin and a few from subdomain 3.

The third contact strengthens the second one by recruitment of additional loops from the upper domain of the 50-kDa fragment of S1 and subdomain 2 of actin. This interaction follows the cleft closure between the lower and upper domains of 50-kDa fragment of S1.

The formation of tightly bound state from the weakly bound state occurs with the second and third contacts. Strong binding is necessary for the power stroke.

Arginine 405, participating in the third contact, is a conserved amino acid residue in all studied myosins. It is functionally important; namely, it was demonstrated that mutation of arginine 405 to glutamine is one cause of familial hypertrophic cardiomyopathy.

These three contacts involve a single actin subunit and define the primary site of actin-myosin interaction. In addition, the model building brings a second region of S1 into the proximity of the neighboring actin monomer, one helix turn below. The main contribution to this secondary interaction with actin comes from a loop that corresponds to residues 567-578 in the S1 sequence.

Remarkable functional conservation is found in the way in which actin and myosin isoforms from different species interact. Such findings suggest a constancy of the structure of the molecular contacts at the actomyosin interface.

Lever arm model: The current hypothesis for the mechanism of force generation by actin-myosin-ATP postulates the rotation of the lever arm as the primary mechanical component of the power stroke. The lever arm is a 85Å-long a helix, to which the regulatory and essential myosin light chains are bound. During the ATP-binding and subsequent ATP-hydrolysis, the active site area of subfragment 1 undergoes subtle conformational changes and as a result the lever arm moves through 11 nm along the actin helix axis (Figs. AM1a and AM1b). Now, it seems likely that the myosin-head power stroke works by such a mechanism. The rotation of the lever arm (Dominguez et al., 1998) that is thought to occur during force production is illustrated in Fig. AM1c.

Fig. AM1a. Illustration of the lever arm's position in the pre-power-stroke state of actomyosin (From Geeves and Holmes, with permission from the Annual Review of Biochemistry, vol. 68,1999, by Annual Reviews, http://www.AnnualReviews.org). Five different actin monomers in two strands of the actin double helix are shown right, a myosin cross-bridge (S1 in Fig. M9) is shown left.

Fig. AM1b. Illustration of the lever arm's position in the post-power-stroke state of actomyosin (From Geeves and Holmes, with permission from the Annual Review of Biochemistry, vol. 68,1999, by Annual Reviews, http://www.AnnualReviews.org).

Fig. AM1c. Illustration of the rotation of the lever arm during force production (From Highsmith, reprinted with permission from Biochemistry 38, 791-797,1999, Copyright 1999, American Chemical Society).

X-ray diffraction and electron microscopy of muscle: The diffraction studies have the advantage of dealing with intact, living muscle under physiological conditions but the interpretation of the diffraction pattern is not straightforward. Electron microscopic pictures visualize the structure but the preparation procedure may produce artifacts.

Because of its striation, skeletal muscle behaves like a diffraction grating with lines spaced apart at a regular interval equal to the sarcomere length, S. Fig. AM2 illustrates the scheme, from which the basis of diffraction may be understood.

Fig. AM2. Theory of X-ray diffraction of muscle (From Wilkie, 1968).

Points A and C represent the Z lines, thus AC=S, the sarcomere length. As the sine waves reach the points A and C they can be regarded as a new light source, scattering light in all directions. Along some directions the scattered waves will be in phase, so the waves reinforce one another and the light is bright. The condition for the first bright fringe is that the path difference AB should be one full wavelength of the light. Thus AB = l. Then from the triangle ABC, sin q = l /S. The wavelength, l, is known, the angle, q, can be experimentally measured and S can be calculated.

As one can measure sarcomere length with the diffraction method, one can also measure the length of the thick and thin filaments.

Hugh Huxley pioneered in obtaining X-ray diffraction pictures from live frog sartorius muscles (Huxley and Brown, 1967, Huxley 1968), one of his earliest results are shown on Fig. AM3. The longitudinal (meridional) dots arise from the myosin filaments and the horizontal (equatorial) lines from the crossbridges. The numbers represent the reflection of the X-ray beam from the ultrastructure. (The white rectangle in the center and the diagonal line are artifacts). Fig. AM4 shows the model deduced from the X-ray diagram. The diffraction pattern of layer-line reflections corresponds to a repeat of 429 Å and arises from the helical arrangement of crossbridges on the myosin filaments. There are pairs of crossbridges on either side of the thick filaments at 143 Å intervals, with each pair rotated by 120o with respect to each neighbor.

Fig. AM3. X-ray diffraction pattern of live frog muscle (From Wilkie, 1968).

 

Fig. AM4. Schematic diagram for arrangement of crossbridges in X-ray diagram of frog muscle (From Needham, 1971).

 

Fig. AM5 compares the equatorial X-ray patterns from live muscle with muscle in rigor. Note the changes in the intensities of the 10 and 11 crystallographic directions. In the resting muscle most of the intensity is in 10 and only very little in 11, whereas in the rigor muscle the intensity of 11 is increased greatly. This is explained by the separate entities of myosin and actin filaments in the resting muscle, whereas in the rigor muscle a large part of the myosin heads is attached to actin.

Fig. AM5. Low-angle equatorial X-ray patterns from rabbit psoas muscle: (top) live; (bottom) in rigor. (Reprinted from J.Mol.Biol. vol.37. Huxley, H.E."Structural difference between resting and rigor muscle; evidence from intensity changes in the low-angle equatorial X-ray diagram", pp 507- 520, 1998, by permission of the publisher, Academic Press)..

The pattern shows the 10 and 11 reflections from the hexagonal lattice of myosin and actin filaments.

 

Rapid freezing electron microscopy detected structural changes in the crossbridges accompanying force generation (Hirose et al., 1994). The structure changed dramatically between relaxed, rigor and with time after initiating contraction. The results supported the models of muscle contraction that have attributed force generation to structural changes in attached crossbridges. With further development of the technique, three-dimensional electron miicroscope tomography was able to visualize directly motor actions of myosin during contraction of insect flight muscle (Taylor et al., 1999).

In vitro motility assay: Sheetz and Spudich (1983) have introduced the in vitro motility assay for studying actin-myosin interaction at the molecular level. In the motility assay either myosin-coated fluorescent beads are observed moving unidirectionally along organized actin filament arrays or single fluorescent-labeled actin filaments are visualized moving over a myosin-coated surface. Movement is initiated by the addition of ATP.

Experiments with the motility assay have shown that myosin determines the actin filament velocity: Pure populations and mixtures of fast skeletal myosin, V1 and V3 cardiac myosin, phosphorylated and unphosphorylated smooth muscle myosin were studied and the movement of fluorescent-labeled actin filaments over myosin coated surfaces was observed. The actin filament velocities were proportional to the actin-activated ATPase activities of the myosins. Furthermore, when the same type of myosin, Dictyostelium, was used as the coated surface, two different actins, Dictyostelium and skeletal muscle, moved with the same velocity, about 2 mm/s, but the same actins moved about 5 mm/s when skeletal muscle myosin was used as the surface. Thus, under all conditions the myosin moiety has controlled the speed of movement.

An "optical tweezers" technique (Fig. AM6) was developed in Spudich's laboratory to measure both the force and step size of single myosin molecules (Finer et al., 1994). Single force transients of 3-4 pN were obtained for movement of 11 nm.

Fig. AM6. IIlustration of two optical traps that are focused on beads attached to a single actin filament, which is held near a single heavy meromyosin (HMM) molecule. The filament is pulled taut and lowered onto a silica bead that is firmly fixed to a microscope coverslip and sparsely coated with HMM (From Finer et al. , reproduced with permission from Nature 368, 113-119,1994, http:// www.nature.com).

References

Dominguez, R., Freyzon, Y. Trybus, K.M., and Cohen, C. (1998). Crystal structure of vertebrate smooth muscle myosin motor domain and its complex with the essential light chain: visualizatioin of the pre-power stroke state. Cell, 94, 559-571.

Finer, J.T., Simmons, R.M. and Spudich, J.A. (1994). Single myosin molecule mechanics: piconewton force and nanometre steps. Nature, 368, 113-119.

Geeves, M.A. and Holmes, K.C. (1999). Structural mechanism of muscle contraction. Annu. Rev. Biochem. 68, 687-728.

Highsmith, S. (1999). Lever arm model for force generation by actin-myosin-ATP. Biochemistry, 38, 791-797.

Hirose, K., Franzini-Armstrong, C., Goldman, Y.E., and Murray, J.M. (1994). Structural changes in muscle crossbridges accompanying force generation. J. Cell. Biol. 127, 763-768.

Huxley, H.E. (1968). Structural difference between resting and rigor muscle; evidence from intensity changes in the low-angle equatorial X-ray diagram. J. Mol. Biol. 37, 507-520.

Huxley, H.E. and Brown W. (1967). The low angle X-ray diagram of vertebrate striated muscle and its behavior during contraction and rigor. J. Mol. Biol. 30, 383-434.

Needham, D. M. (1971). Machina Carnis. Cambridge University Press

Rayment, I. and Holden, H.M. (1995). In, Molecular Aspects of Cell Biology (R. H. Garrett and C.M. Grisham Eds), Saunders College Publishing.

Sheetz, M.P. and Spudich, J.A. (1983). Movement of myosin-coated fluorescent beads on actin cables in vitro. Nature, 303, 31-35.

Taylor, K.A., Schmitz, H., Reedy, M.C., Goldman, Y.E., Franzinini-Armstrong, C., Sasaki, H., Tregear, R.T., Poole, K., Lucaveche, C., Edwards, R.J., Chen, L.F., Winkler, H., and Reedy, M. K. (1999). Tomographic 3D reconstruction of quick frozen, Ca2+ -activated contracting insect flight muscle. Cell, 99, 421-431.

Wilkie, D.R. (1968). Muscle. St. Martin Press.


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