Advertisement

Myosin: Cellular Molecular Motor

Chapter

Abstract

Myosins are a superfamily of ATP-dependent, actin-based motor proteins. In addition to muscle contraction, myosins are involved in a wide range of motility functions in cells, including transport of intracellular cargo. Typically, myosin molecules are composed of a heavy chain having a tail, hinge, and head domain and light chain present near the myosin head that modulates calcium-dependent transduction of force by myosin. The myosin head has both actin binding and ATP binding and ATPase activity. In resting state, myosin is bound to actin. When ATP binds to the myosin head, the head dissociates from the actin filament. Hydrolysis of bound ATP to ADP and the release of phosphate establish the rigor state of myosin (reestablishes myosin–actin interaction), generating force in the process to walk along the actin filament. Binding of a new ATP molecule to the myosin head releases myosin from actin to repeat the cycle. The structure and function of myosin are globally conserved.

Keywords

Myosin ATP-fueled motor proteins Actin-based motility 

References

  1. 1.
    Kühne, W. (1859). Untersuchungen über bewegungen und veränderungen der kontraktileu substanzen. Archiv für Anatomie. Physiologie und wissenschaftliche Medicin, 1859, 748–783.Google Scholar
  2. 2.
    Engelhardt, W. A., & Ljubimowa, M. N. (1939). Myosine and adenosine triphosphate. Nature, 144, 668–669.Google Scholar
  3. 3.
    Banga, I., & Szent-Gyorgyi, A. (1941). Preparation and properties of myosin A and B. In Studies from the Institute of Medical Chemistry University of Szeged, edited by Szent-Gyorgyi A (Vol. I, pp. 5–15). New York: Karger.Google Scholar
  4. 4.
    Banga, I., & Szent-Gyorgyi, A. (1943). The influence of salts on the phosphatase action of myosin. In A. Szent-Gyorgyi (Ed.), Studies from the Institute of Medical Chemistry University of Szeged (Vol. III, pp. 72–75). New York: Karger.Google Scholar
  5. 5.
    Straub, F. B. (1942). Actin. In A. Szent-Gyorgyi (Ed.), Studies from the Institute of Medical Chemistry University of Szeged (Vol. II, pp. 3–15). New York: Karger.Google Scholar
  6. 6.
    Straub, F. B. (1943). Actin II. In Studies from the Institute of Medical Chemistry University of Szeged, edited by Szent-Gyorgyi A (Vol. III, pp. 23–37). New York: Karger.Google Scholar
  7. 7.
    Huxley, A. F., & Niedergerke, R. (1954). Structural changes in muscle during contraction: Interference microscopy of living muscle fibres. Nature, 173, 971–973.PubMedGoogle Scholar
  8. 8.
    Huxley, H. E., & Hanson, J. (1954). Changes in the cross-striations of muscle during contraction and stretch and their structural interpretation. Nature, 173, 973–976.PubMedGoogle Scholar
  9. 9.
    Huxley, H. E. (1963). Electron microscope studies on the structure of natural and synthetic protein filaments from striated muscle. Journal of Molecular Biology, 7, 281–308.PubMedGoogle Scholar
  10. 10.
    Slayter, H. S., & Lowey, S. (1967). Substructure of the myosin molecule as visualized by electron micros- copy. Proceedings of the National Academy of Sciences, 58, 1611–1618.Google Scholar
  11. 11.
    Lowey, S., Slayter, H. S., Weeds, A. G., & Baker, H. (1969). Substructure of the myosin molecule. I. Subfragments of myosin by enzymic degradation. Journal of Molecular Biology, 42, 1–29.PubMedGoogle Scholar
  12. 12.
    Kendrick-Jones, J., Lehman, W., & Szent-Gyorgyi, A. G. (1970). Regulation in molluscan muscles. Journal of Molecular Biology, 54, 313–326.PubMedGoogle Scholar
  13. 13.
    Greaser, M. L., & Gergely, J. (1971). Reconstitution of troponin activity from three protein components. The Journal of Biological Chemistry, 246, 4226–4233.PubMedGoogle Scholar
  14. 14.
    Huxley, A. F., & Simmons, R. M. (1971). Proposed mechanism of force generation in striated muscle. Nature, 233, 533–538.PubMedGoogle Scholar
  15. 15.
    Lymn, R. W., & Taylor, E. W. (1971). Mechanism of adenosine triphosphate hydrolysis by actomyosin. Biochemistry, 10, 4617–4624.PubMedGoogle Scholar
  16. 16.
    Bremel, R. D., & Weber, A. (1972). Cooperation within actin filament in vertebrate skeletal muscle. Nature: New Biology, 238, 97–101.Google Scholar
  17. 17.
    Haselgrove, J. C., & Huxley, H. E. (1973). X-ray evidence for radial cross-bridge movement and for the sliding filament model in actively contracting skeletal muscle. Journal of Molecular Biology, 77, 549–568.PubMedGoogle Scholar
  18. 18.
    Bagshaw, C. R., & Trentham, D. R. (1974). The characterization of myosin-product complexes and of product-release steps during the magnesium ion-dependent adenosine triphosphatase reaction. The Biochemical Journal, 141, 331–349.PubMedPubMedCentralGoogle Scholar
  19. 19.
    Hibberd, M. G., Dantzig, J. A., Trentham, D. R., & Goldman, Y. E. (1985). Phosphate release and force generation in skeletal muscle fibers. Science, 228, 1317–1319.PubMedGoogle Scholar
  20. 20.
    Kron, S. J., & Spudich, J. A. (1986). Fluorescent actin filaments move on myosin fixed to a glass surface. Proceedings of the National Academy of Sciences, 83, 6272–6276.Google Scholar
  21. 21.
    Toyoshima, Y. Y., Kron, S. J., McNally, E. M., Niebling, K. R., Toyoshima, C., & Spudich, J. A. (1987). Myosin subfragment-1 is sufficient to move actin filaments in vitro. Nature, 328, 536–539.PubMedGoogle Scholar
  22. 22.
    Laha, S., Naik, A. R., Kuhn, E. R., Alvarez, M., Sujkowski, A., Wessels, R. J., & Jena, B. P. (2017). Nanothermometry measure of muscle efficiency. ACS Nano Letters, 17, 1262–1268.Google Scholar
  23. 23.
    Kuhn, E. R., Naik, A. R., Lewis, B. E., Kokotovich, K. M., Li, M., Stemmler, T. M., et al. (2018). Nanothermometry reveals calcium-induced remodeling of myosin. ACS Nano Letters, 18, 7021–7029.Google Scholar
  24. 24.
    Kronert, W. A., Bell, K. M., Viswanathan, M. C., Melkani, G. C., Trujillo, A. S., Huang, A., et al. (2018). Prolonged cross-bridge binding triggers muscle dysfunction in a fly model of myosin-based hypertrophic cardiomyopathy. eLife, 7, e38e064.Google Scholar
  25. 25.
    Li, M., Ogilvie, H., Ochala, J., Konstantin, A., Iwamoto, H., Yagi, N., et al. (2015). Aberrant post-translational modifications compromise human myosin motor function in old age. Aging Cell, 14, 228–235.PubMedPubMedCentralGoogle Scholar
  26. 26.
    Llsano-Diez, M., Renaud, G., Andersson, M., Marrero, H. G., Cacciani, N., Engquist, H., Corpeno, R., Artemenko, K., Bergquist, J., & Larsson, L. (2012). Mechanism underlying ICU muscle wasting and effects of passive mechanical loading. Critical Care, 16, R209.Google Scholar
  27. 27.
    Llsano-Diez, M., Gustafsin, A.-M., Olsson, C., Goransson, H., & Larsson, L. (2011). Muscle wasting and the temporal gene expression pattern in a novel rat intensive care unit model. BMC Genomics, 12, 602.Google Scholar
  28. 28.
    Norman, H., Nordquist, J., Andersson, P., Ansved, T., Tang, X., Dworkin, B., et al. (2006). Impact of post-synaptic block of neuromuscular transmission, muscle unloading and mechanical ventilation on skeletal muscle protein and mRNA expression. Pflügers Archiv, 453, 53–66.PubMedGoogle Scholar
  29. 29.
    Larsson, L. (2007). Experimental animal models of muscle wasting in intensive care unit patients. Critical Care Medicine, 35(9 Suppl), S484–S487.PubMedGoogle Scholar
  30. 30.
    Ochala, J., Gustafson, A.-M., Li, M., Aare, S., Qaisar, R., Llano Diez, M., et al. (2011). Preferential skeletal muscle myosin loss in response to mechanical silencing in a novel rat intensive care unit model: underlying mechanisms. The Journal of Physiology, 589(8), 2007–2026.PubMedPubMedCentralGoogle Scholar
  31. 31.
    Akkad, H., Corpeno, R., & Larsson, L. (2014). Masseter muscle myofibrillar protein synthesis and degradation in an experimental critical illness myopathy model. Plos-One., 9(4), e92622.PubMedPubMedCentralGoogle Scholar
  32. 32.
    Pette, D., & Staron, R. S. (2000). Myosin isoforms, muscle fiber types, and transitions. Microscopy Research and Technique, 50(6), 500–509.PubMedGoogle Scholar
  33. 33.
    MacFarlane, I. A., & Rosenthal, F. D. (1977). Severe myopathy after status asthmaticus. Lancet, 2(8038), 615.PubMedGoogle Scholar
  34. 34.
    Lacomis, D., Zochodne, D. W., & Bird, S. J. (2000). Critical illness myopathy. Muscle & Nerve, 23(12), 1785–1788.Google Scholar
  35. 35.
    Phillips, S. M., Glover, E. I., & Rennie, M. J. (2009). Alterations of protein turnover underlying disuse atrophy in human skeletal muscle. Journal of Applied Physiology, 107(3), 645–654.PubMedGoogle Scholar
  36. 36.
    Cohen, S., Brault, J. J., Gygi, S. P., et al. (2009). During muscle atrophy, thick, but not thin, filament components are degraded by MuRF1-dependent ubiquitylation. The Journal of Cell Biology, 185(6), 1083–1095.PubMedPubMedCentralGoogle Scholar
  37. 37.
    Lecker, S. H., Jagoe, R. T., Gilbert, A., et al. (2004). Multiple types of skeletal muscle atrophy involve a common program of changes in gene expression. The FASEB Journal, 18(1), 39–51.PubMedGoogle Scholar
  38. 38.
    Sacheck, J. M., Hyatt, J. P., Raffaello, A., et al. (2007). Rapid disuse and denervation atrophy involve transcriptional changes similar to those of muscle wasting during systemic diseases. The FASEB Journal, 21(1), 140–155.PubMedGoogle Scholar
  39. 39.
    Sandri, M. (2010). Autophagy in health and disease. 3. Involvement of autophagy in muscle atrophy. American Journal of Physiology. Cell Physiology, 298(6), C1291–C1297.PubMedGoogle Scholar
  40. 40.
    Stevenson, E., Giresi, P., Koncarevic, A., & Kandarian, S. (2003). Global analysis of gene expression patterns during disuse atrophy in rat skeletal muscle. Journal of Physiology-London., 2003, 33–48.Google Scholar
  41. 41.
    Sasa, T., Sairyo, K., Yoshida, N., et al. (2004). Continuous muscle stretch prevents disuse muscle atrophy and deterioration of its oxidative capacity in rat tail-suspension models. American Journal of Physical Medicine & Rehabilitation, 83(11), 851–856.Google Scholar
  42. 42.
    Sandri, M., Lin, J., Handschin, C., Yang, W., Arany, Z. P., Lecker, S. H., et al. (2006). PGC-1alpha protects skeletal muscle from atrophy by suppressing FoxO3 action and atrophy- specific gene transcription. Proceedings of the National Academy of Sciences of the United States of America, 103, 16260–16265.PubMedPubMedCentralGoogle Scholar
  43. 43.
    Ruas, J. L., White, J. P., Rao, R. R., Kleiner, S., Brannan, K. T., Harrison, B. C., et al. (2012). A PGC-1alpha isoform induced by resistance training regulates skeletal muscle hypertrophy. Cell, 151, 1319–1331.PubMedPubMedCentralGoogle Scholar
  44. 44.
    Resnicow, R., Deacon, J. C., Warrick, H. M., Spudich, J. A., & Leinwand, L. A. (2010). Functional diversity among a family of human skeletal muscle myosin motors. Proceedings of the National Academy of Sciences of the United States of America, 107, 1053–1058.PubMedGoogle Scholar
  45. 45.
    Schiaffino, S., & Reggiani, C. (1994). Myosin isoforms in mammalian skeletal muscle. Journal of Applied Physiology, 77, 493–501.PubMedGoogle Scholar
  46. 46.
    Schiaffino, S., & Reggiani, C. (1996). Molecular diversity of myofibrillar proteins: Gene regulation and functional significance. Physiological Reviews, 76, 371–423.PubMedGoogle Scholar
  47. 47.
    Pette, D., & Staron, R. S. (1997). Mammalian skeletal muscle fiber type transitions. International Review of Cytology, 170, 143–223.PubMedGoogle Scholar
  48. 48.
    Staron, R. S., & Pette, D. (1987). Nonuniform myosin expression along single fibers of chronically stimulated and contralateral rabbit tibialis anterior muscles. Pflügers Archiv, 409, 67–73.PubMedGoogle Scholar
  49. 49.
    Peuker, H., & Pette, D. (1997). Quantitative analyses of myosin heavy-chain mRNA and protein isoforms in single fibers reveal a pronounced fiber heterogeneity in normal rabbit muscles. European Journal of Biochemistry, 247, 30–36.PubMedGoogle Scholar
  50. 50.
    Andersen, J. L., & Schiaffino, S. (1997). Mismatch between myosin heavy chain mRNA and protein distribution in human skeletal muscle fibers. American Journal of Physiology-Cell Physiology, 272, C1881–1889. 37.Google Scholar
  51. 51.
    Eizema, K., et al. (2003). Differential expression of equine myosin heavy-chain mRNA and protein isoforms in a limb muscle. The Journal of Histochemistry and Cytochemistry, 51, 1207–1216.PubMedGoogle Scholar
  52. 52.
    Smith, K., & Rennie, M. J. (1996). The measurement of tissue protein turnover. Baillière’s Clinical Endocrinology and Metabolism, 10, 469–495.PubMedGoogle Scholar
  53. 53.
    Mann, M., & Jensen, O. (2003). Proteomic analysis of post-translational modifications. Nature Biotechnology, 21, 255–261.PubMedGoogle Scholar
  54. 54.
    Kim, W., Bennett, E. J., Huttlin, E. L., Guo, A., Li, J., Possemato, A., et al. (2011). Systematic and quantitative assessment of the ubiquitin-modified proteome. Molecular Cell, 44, 325–340.PubMedPubMedCentralGoogle Scholar
  55. 55.
    Edwards, A. V. G., Cordwell, S. J., & White, M. Y. (2011). Phosphoproteomic pro ling of the myocyte. Circulation. Cardiovascular Genetics, 4, 575.PubMedGoogle Scholar
  56. 56.
    Hennet, T. (2012). Diseases of glycosylation beyond classical congenital disorders of glycosylation. Biochimica et Biophysica Acta, 1820, 1306–1317.PubMedGoogle Scholar
  57. 57.
    Li, T., Evdokimov, E., Shen, R.-F., Chao, C.-C., Tekle, E., Wang, T., Stadtman, E. R., Yang, D. C. H., & Chock, P. B. (2004). Sumoylation of heterogeneous nuclear ribonucleoproteins, zinc finger proteins, and nuclear pore complex proteins: A proteomic analysis. Proceedings of the National Academy of Sciences of the United States of America, 101, 8551–8556.PubMedPubMedCentralGoogle Scholar
  58. 58.
    Matafora, V., D’Amato, A., Mori, S., Blasi, F., & Bachi, A. (2009). Proteomics analysis of nucleolar SUMO-1 target proteins upon proteasome inhibition. Molecular & Cellular Proteomics, 8, 2243–2255.Google Scholar
  59. 59.
    Flick, K., & Kaiser, P. (2009). Proteomic revelation: SUMO changes partners when the heat is on. Science Signaling, 2, pe45.PubMedPubMedCentralGoogle Scholar
  60. 60.
    Tatham, M. H., Rodriguez, M. S., Xirodimas, D. P., & Hay, R. T. (2009). Detection of protein SUMOylation in vivo. Nature Protocols, 4, 1363–1371.PubMedGoogle Scholar
  61. 61.
    Galisson, F., Mahrouche, L., Courcelles, M., Bonneil, E., Meloche, S., Chelbi-Alix, M. K., & Thibault, P. (2011). A novel proteomics approach to identify SUMOylated proteins and their modification sites in human cells. Molecular & Cellular Proteomics, 10, M110.004796.Google Scholar
  62. 62.
    Norman, H., Nordquist, J., Andersson, P., Ansved, T., Tang, X., Dworkin, B., & Larsson, L. (2006). Impact of post-synaptic block of neuromuscular transmission, muscle unloading and mechanical ventilation on skeletal muscle protein and mRNA expression. Pflügers Archiv, 453, 53–66.PubMedGoogle Scholar
  63. 63.
    Lee, S. Y., Ahn, S., Kim, Y. J., Ji, M. J., Kim, K. M., Choi, S. H., et al. (2018). Comparison between dual-energy X-ray absorptiometry and bioelectrical impedance analyses for accuracy in measuring whole body muscle mass and appendicular skeletal muscle mass. Nutrients, 10, 738.PubMedCentralGoogle Scholar
  64. 64.
    Mitsiopoulos, N., Baumgartner, R. N., Heymsfield, S. B., Lyons, W., Gallagher, D., & Ross, R. (1998). Cadaver validation of skeletal muscle measurement by magnetic resonance imaging and computerized tomography. Journal of Applied Physiology, 85, 115–122.PubMedGoogle Scholar
  65. 65.
    Reeves, N. D., Maganaris, C. N., & Narici, M. V. (2004). Ultrasonographic assessment of human skeletal muscle size. European Journal of Applied Physiology, 91, 116–118.PubMedGoogle Scholar
  66. 66.
    Mourtzakis, M., Parry, S., Connolly, B., & Puthucheary, Z. (2017). Skeletal muscle ultrasound in critical care: A tool in need of translation. Annals of the American Thoracic Society, 14, 1495–1503.PubMedPubMedCentralGoogle Scholar
  67. 67.
    Mercuri, E., Pichiecchio, A., Allsop, J., Messina, S., Pane, M., & Muntoni, F. (2007). Muscle MRI in inherited neuromuscular disorders: Past, present, and future. Journal of Magnetic Resonance Imaging, 25(2), 433–440.PubMedGoogle Scholar
  68. 68.
    Mercuri, E., Jungbluth, H., & Muntoni, F. (2005). Muscle imaging in clinical practice: Diagnostic value of muscle magnetic resonance imaging in inherited neuromuscular disorders. Current Opinion in Neurology, 18(5), 526–537.PubMedGoogle Scholar
  69. 69.
    Zaidman, C. M., Holland, M. R., Anderson, C. C., & Pestronk, A. (2008). Calibrated quantitative ultrasound imaging of skeletal muscle using backscatter analysis. Muscle & Nerve, 38(1), 893–898.Google Scholar
  70. 70.
    Lai, C. H., Melli, G., Chang, Y. J., Skolasky, R. L., Corse, A. M., Wagner, K. R., & Cornblath, D. R. (2010). Open muscle biopsy in suspected myopathy: Diagnostic yield and clinical utility. European Journal of Neurology, 17(1), 136–142.PubMedGoogle Scholar
  71. 71.
    Collins, M. P., Mendell, J. R., Periquet, M. I., Sahenk, Z., Amato, A. A., Gronseth, G. S., et al. (2000). Superficial peroneal nerve/peroneus brevis muscle biopsy in vasculitic neuropathy. Neurology, 55(5), 636–643.PubMedGoogle Scholar
  72. 72.
    Tarnopolsky, M. A., Pearce, E., Smith, K., & Lach, B. (2011). Suction-modified Bergström muscle biopsy technique: Experience with 13,500 procedures. Muscle & Nerve, 43(5), 717–725.Google Scholar
  73. 73.
    Giagnacovo, M., Cardani, R., Meola, G., Pellicciari, C., & Malatesta, M. (2010). Routinely frozen biopsies of human skeletal muscle are suitable for morphological and immunocytochemical analyses at transmission electron microscopy. European Journal of Histochemistry, 54(3), e31.PubMedGoogle Scholar
  74. 74.
    Bossen, E. (2000). In R. Wortmann (Ed.), Muscle biopsy in disease of skeletal muscle (pp. 333–348). Philadelphia, PA: Lippincott Williams and Wilkins.Google Scholar
  75. 75.
    Dalakas, M. C. (2002). Muscle biopsy findings in inflammatory myopathies. Rheumatic Diseases Clinics of North America, 28(4), 779–798.PubMedGoogle Scholar
  76. 76.
    Scola, R. H., Pereira, E. R., Lorenzoni, P. J., & Werneck, L. C. (2007). Toxic myopathies: Muscle biopsy features. Arquivos de Neuro-Psiquiatria, 65(1), 82–86.PubMedGoogle Scholar
  77. 77.
    Rifai, Z., Welle, S., Kamp, C., & Thornton, C. A. (1995). Ragged red fibers in normal aging and inflammatory myopathy. Annals of Neurology, 37(1), 24–29.PubMedGoogle Scholar
  78. 78.
    Jungbluth, H., Sewry, C. A., & Muntoni, F. (2011). Core myopathies. Seminars in Pediatric Neurology, 18(4), 239–249.PubMedGoogle Scholar
  79. 79.
    Huizing, M., & Krasnewich, D. M. (2009). Hereditary inclusion body myopathy: A decade of progress. Biochimica et Biophysica Acta, 1792(9), 881–887.PubMedPubMedCentralGoogle Scholar
  80. 80.
    Pernal, S.P., Liyanaarachchi, A., Gatti, D.L., Formosa, B., Pulvender, R., Kuhn, E.R., et al. (2019) Differential expansion microscopy. bioRxiv. doi: https://doi.org/10.1101/699579.
  81. 81.
    Gatti, D.L., Arslanturk, S., Lal, S., Jena, B.P. (2019) Deep learning strategies for differential expansion microscopy. bioRxiv. doi: https://doi.org/10.1101/743682.

Copyright information

© Springer Nature Switzerland AG 2020

Authors and Affiliations

  1. 1.Department of PhysiologyWayne State University School of MedicineDetroitUSA

Personalised recommendations