Forcing cells into shape: the mechanics of actomyosin contractility

Journal name:
Nature Reviews Molecular Cell Biology
Volume:
16,
Pages:
486–498
Year published:
DOI:
doi:10.1038/nrm4012
Published online

Abstract

Actomyosin-mediated contractility is a highly conserved mechanism for generating mechanical stress in animal cells and underlies muscle contraction, cell migration, cell division and tissue morphogenesis. Whereas actomyosin-mediated contractility in striated muscle is well understood, the regulation of such contractility in non-muscle and smooth muscle cells is less certain. Our increased understanding of the mechanics of actomyosin arrays that lack sarcomeric organization has revealed novel modes of regulation and force transmission. This work also provides an example of how diverse mechanical behaviours at cellular scales can arise from common molecular components, underscoring the need for experiments and theories to bridge the molecular to cellular length scales.

At a glance

Figures

  1. Types of contractile deformations generated by cells and tissues.
    Figure 1: Types of contractile deformations generated by cells and tissues.

    a | Isotropic contraction, which is performed by platelets, is uniform around the cell perimeter and induces a uniform change in shape and in the force generated. b | Anisotropic contraction, which is performed by striated and smooth muscle cells, induces contraction and force generation along one axis. c–e | In cytokinesis and cell migration, contractile stresses are spatially localized in a particular region of the cell to generate large deformations in cytokinesis (panel c), during symmetry breaking in migrating cells (panel d) and during tail retraction in migrating cells (panel e). f | Immunofluorescence images of several adherent cell types stained for actin, myosin II and α-actinin, including human platelets, striated muscle from a rat heart, smooth muscle from a human airway and mouse NIH 3T3 fibroblasts. Insets are a magnification of the corresponding boxed region and highlight the actomyosin organization within the cell. Myosin II was visualized using an antibody against phosphorylated myosin light chain, α-actinin was visualized via direct antibody staining and actin was visualized via phalloidin staining. Image of striated muscle cell courtesy of B. Hissa, University of Chicago, Illinois, USA, and image of smooth muscle cell courtesy of Y. Beckham, University of Chicago, Illinois, USA.

  2. Contractility in sarcomeres.
    Figure 2: Contractility in sarcomeres.

    a | Filamentous (F)-actin has a barbed end and a pointed end (indicated by the 'open' and 'closed' direction, respectively, of the depicted actin chevrons that make up the polymer), and it can associate with globular (G)-actin from a pool of monomers or add G-actin back to this pool, as indicated by the arrows. Higher association rates of monomeric actin to the barbed end are indicated by a larger arrow. Bipolar myosin filaments with a central bare zone that lacks motor heads are assembled from myosin II dimers. b | Myosin II filaments drive the translocation of F-actin filaments towards their barbed ends with a characteristic force (F) and gliding velocity (v) relationship. This can result in the contraction (left) or extension (right) of two bound actin filaments, depending on the location of myosin II with respect to the middle of these filaments. c | Actomyosin organization within sarcomeres. Here myosin filaments are segregated towards the F-actin pointed ends, and F-actin barbed ends are localized at Z-bands, which contain numerous regulatory proteins, including α-actinin crosslinkers. The initial and final contractile unit length are indicated by li and lf, respectively, in the initial (i) and final (f) states. Black arrows in the initial state indicate the direction of F-actin translocation. The contractile unit size is set by the sarcomere geometry, with the bundle shortening velocity (V) equal to the number of contractile units (N) times the myosin gliding velocity (v) such that V = Nv. The reduction in sarcomere length between the initial state and the final state arises from increased overlap between the F-actin and the myosin bare zone. The entire bundle length L is determined by the number N of contractile units multiplied by their length (l). Thus, the initial and final bundle lengths are given by Li = Nli and Lf = Nlf respectively.

  3. Contractility in disordered actomyosin bundles.
    Figure 3: Contractility in disordered actomyosin bundles.

    Throughout the figure the initial (i) and final (f) configurations are indicated, and the initial and final contractile unit length is indicated by li and lf, respectively. Actin chevrons indicate the direction of the actin; the actin barbed end and actin pointed end are depicted by the 'open' and 'closed' direction of the chevrons, respectively. a | In a bundle with disordered actomyosin orientations, myosin II activity results in the internal sorting of F-actin polarity but does not lead to an overall reduction in the average bundle length (the initial and final bundle lengths are equal Lf = Li). b | Quasi-sarcomeric organizations arise in some cytoskeletal assemblies, such as the Schizosaccharomyces pombe contractile ring, in which myosin II and formins are localized to nodes. Formins cluster F-actin barbed ends, thus localizing myosin II activity from neighbouring nodes towards F-actin pointed ends. This is effectively a sarcomere-like geometry and results in contractility over time. c | In a disordered actomyosin bundle similar to that shown in part a but with added crosslinking, myosin II activity generates internal compressive and tensile stresses, which cause the compression or extension of F-actin portions, respectively, depending on the relative position of motors and crosslinks with respect to F-actin barbed ends. If sufficiently large, these internal stresses deform and buckle portions of F-actin, which relieves compressive stress and enables bundle shortening. In this model, the average contractile unit size is the average distance between F-actin buckling events.

  4. Inherent contractility of adherent cells.
    Figure 4: Inherent contractility of adherent cells.

    a | Traction force microscopy measures the distribution and magnitude of traction stresses of adherent cells (indicated in red) exerted through focal adhesions (green ovals) by probing the deformation of the underlying compliant matrix (grey) as measured by fiduciary markers (grey circles). Using this technique, we can calculate the strain in the substrate (u; grey arrows), the traction stresses applied by the cell (T; white arrows) and the amount of work performed to deform the substrate (W). The strain and stress are both vector fields, meaning that at each position these quantities have both a direction and magnitude. The total work is determined by integrating the dot product of the strain and stress vectors over the entire area (dA). b | The traction stress direction and magnitude for NIH 3T3 fibroblast cells of similar areas (~1,600 μm2) plated on a circular (top), oblong (middle) and unpatterned (bottom) surface are shown. The cell area is approximately constant in each of the three conditions, resulting in a similar amount of mechanical work performed on the environment by each of these three cells106. Different cell geometries, however, result in different distributions of stresses (measured in Pa, as indicated) on the surface. c | Numerous experimental groups have found that the strain energy is proportional to the spread area for a wide range of cell types and for multicellular islands. The magnitude of this ratio (that is, slope of the line) is a measure of the characteristic contractility, and thus is cell type dependent. Multicellular islands of epithelial cells scale similarly to those for single cells when the area of the entire island is considered104. d | The ratio of strain energy to spread area shows a characteristic contractility value for different muscle and non-muscle cells.

References

  1. Munjal, A. & Lecuit, T. Actomyosin networks and tissue morphogenesis. Development 141, 17891793 (2014).
  2. Gardel, M. L., Schneider, I. C., Aratyn-Schaus, Y. & Waterman, C. M. Mechanical integration of actin and adhesion dynamics in cell migration. Annu. Rev. Cell Dev. Biol. 26, 315333 (2010).
  3. Vicente-Manzanares, M., Ma, X., Adelstein, R. S. & Horwitz, A. R. Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat. Rev. Mol. Cell Biol. 10, 778790 (2009).
  4. Salbreux, G., Charras, G. & Paluch, E. Actin cortex mechanics and cellular morphogenesis. Trends Cell Biol. 10, 536545 (2012).
  5. Green, R. A., Paluch, E. & Oegema, K. Cytokinesis in animal cells. Annu. Rev. Cell Dev. Biol. 28, 2958 (2012).
  6. Pinto, I. M. et al. Actin depolymerization drives actomyosin ring contraction during budding yeast cytokinesis. Dev. Cell 22, 12471260 (2012).
  7. Murrell, M. P. et al. Liposome adhesion generates traction stress. Nat. Phys. 10, 163169 (2014).
  8. Stroka, K. M. et al. Water permeation drives tumor cell migration in confined microenvironments. Cell 157, 611623 (2014).
  9. Levayer, R. & Lecuit, T. Biomechanical regulation of contractility: spatial control and dynamics. Trends Cell Biol. 22, 6181 (2012).
  10. Lecuit, T., Lenne, P.-F. & Munro, E. Force generation, transmission, and integration during cell and tissue morphogenesis. Ann. Rev. Cell Dev. Bio 27, 157184 (2011).
  11. Gordon, A. M., Homsher, E. & Regnier, M. Regulation of contraction in striated muscle. Physiol. Rev. 80, 853924 (2000).
  12. Huxley, H. E. Fifty years of muscle and the sliding filament hypothesis. Eur. J. Biochem. 271, 14031415 (2004).
  13. Steinmetz, P. R. H. et al. Independent evolution of striated muscles in cnidarians and bilaterians. Nature 487, 231234 (2012).
  14. Niederman, R. & Pollard, T. D. Human platelet myosin. II. In vitro assembly and structure of myosin filaments. J. Cell Biol. 67, 7292 (1975).
  15. Pollard, T. D. Structure and polymerization of Acanthamoeba myosin-II filaments. J. Cell Biol. 95, 816825 (1982).
  16. Skubiszak, L. & Kowalczyk, L. Myosin molecule packing within the vertebrate skeletal muscle thick filaments. A complete bipolar model. Acta Biochim. Polon. 49, 829840 (2002).
  17. Sobieszek, A. Cross-bridges on self-assembled smooth muscle myosin filaments. J. Mol. Biol. 70, 741744 (1972).
  18. Tonino, P., Simon, M. & Craig, R. Mass determination of native smooth muscle myosin filaments by scanning transmission electron microscopy. J. Mol. Biol. 318, 9991007 (2002).
  19. Huxley, H. E. X-ray analysis and the problem of muscle. Proc. R. Soc. Lond. B 141, 5962 (1953).
  20. Huxley, H. E. The double array of filaments in cross-striated muscle. J. Biophys. Biochem. Cytol. 3, 631648 (1957).
  21. Huxley, A. F. Muscle structure and theories of contraction. Prog. Biophys. Biophys. Chem. 7, 255318 (1957).
  22. Huxley, A. F. & Niedergerke, R. Structural changes in muscle during contraction: interference microscopy of living muscle fibres. Nature 173, 971973 (1954).
  23. Littlefield, R., Almenar-Queralt, A. & Fowler, V. M. Actin dynamics at pointed ends regulates thin filament length in striated muscle. Nat. Cell Biol. 3, 544551 (2001).
  24. Lavoie, T. L. et al. Disrupting actin–myosin–actin connectivity in airway smooth muscle as a treatment for asthma? Proc. Am. Thorac. Soc. 6, 295300 (2009).
  25. Gunst, S. J. & Zhang, W. Actin cytoskeletal dynamics in smooth muscle: a new paradigm for the regulation of smooth muscle contraction. Am. J. Physiol. Cell Physiol. 295, C576587 (2008).
  26. Verkhovsky, A. B. & Borisy, G. G. Non-sarcomeric mode of myosin II organization in the fibroblast lamellum. J. Cell Biol. 123, 637652 (1993).
  27. Svitkina, T. M., Verkhovsky, A. B., McQuade, K. M. & Borisy, G. G. Analysis of the actin–myosin II system in fish epidermal keratocytes: mechanism of cell body translocation. J. Cell Biol. 139, 397415 (1997).
  28. Aratyn-Schaus, Y., Oakes, P. W. & Gardel, M. L. Dynamic and structural signatures of lamellar actomyosin force generation. Mol. Biol. Cell 22, 13301339 (2011).
  29. Hotulainen, P. & Lappalainen, P. Stress fibers are generated by two distinct actin assembly mechanisms in motile cells. J. Cell Biol. 173, 383394 (2006).
  30. Svitkina, T. M. & Borisy, G. G. Correlative light and electron microscopy of the cytoskeleton of cultured cells. Methods Enzymol. 298, 570592 (1998).
  31. Stricker, J., Beckham, Y., Davidson, M. W. & Gardel, M. L. Myosin II-mediated focal adhesion maturation is tension insensitive. PLoS ONE 8, e70652 (2013).
  32. Oakes, P. W., Beckham, Y., Stricker, J. & Gardel, M. L. Tension is required but not sufficient for focal adhesion maturation without a stress fiber template. J. Cell Bio. 196, 363374 (2012).
  33. Martin, A. C. et al. Integration of contractile forces during tissue invagination. J. Cell Biol. 188, 735749 (2010).
  34. Martin, A. C., Kaschube, M. & Wieschaus, E. F. Pulsed contractions of an actin–myosin network drive apical constriction. Nature 457, 495499 (2009).
  35. He, L., Wang, X., Tang, H. L. & Montell, D. J. Tissue elongation requires oscillating contractions of a basal actomyosin network. Nat. Cell Biol. 12, 11331142 (2010).
  36. Levayer, R. & Lecuit, T. Oscillation and polarity of E-cadherin asymmetries control actomyosin flow patterns during morphogenesis. Dev. Cell 26, 162175 (2013).
  37. Kim, T., Gardel, M. L. & Munro, E. Determinants of fluidlike behavior and effective viscosity in cross-linked actin networks. Biophys. J. 106, 526534 (2014).
  38. Courtemanche, N., Lee, J. Y., Pollard, T. D. & Greene, E. C. Tension modulates actin filament polymerization mediated by formin and profilin. Proc. Natl Acad. Sci. USA 110, 97529757 (2013).
  39. Ferrer, J. M. et al. Measuring molecular rupture forces between single actin filaments and actin-binding proteins. Proc. Natl Acad. Sci. USA 105, 92219226 (2008).
  40. Jégou, A., Carlier, M.-F. & Romet-Lemonne, G. Formin mDia1 senses and generates mechanical forces on actin filaments. Nat. Commun. 4, 1883 (2013).
  41. Wilson, C. A. et al. Myosin II contributes to cell-scale actin network treadmilling through network disassembly. Nature 465, 373377 (2010).
  42. Fritzsche, M. et al. Analysis of turnover dynamics of the submembranous actin cortex. Mol. Biol. Cell 24, 757767 (2013).
  43. Carvalho, A., Desai, A. & Oegema, K. Structural memory in the contractile ring makes the duration of cytokinesis independent of cell size. Cell 137, 926937 (2009).
  44. Luo, W. et al. Analysis of the local organization and dynamics of cellular actin networks. J. Cell Biol. 202, 10571073 (2013).
  45. Lenz, M., Gardel, M. L. & Dinner, A. R. Requirements for contractility in disordered cytoskeletal bundles. New J. Phys. 14, 033037 (2012).
  46. Vavylonis, D. et al. Assembly mechanism of the contractile ring for cytokinesis by fission yeast. Science 319, 97100 (2008).
  47. Kruse, K. & Julicher, F. Actively contracting bundles of polar filaments. Phys. Rev. Lett. 85, 17781781 (2000).
  48. Liverpool, T. B. & Marchetti, M. C. Bridging the microscopic and the hydrodynamic in active filament solutions. Europhys. Lett. 69, 846 (2005).
  49. Tsuda, Y., Yasutake, H., Ishijima, A. & Yanagida, T. Torsional rigidity of single actin filaments and actin–actin bond breaking force under torsion measured directly by in vitro micromanipulation. Proc. Natl Acad. Sci. USA 93, 1293712942 (1996).
  50. McCullough, B. R. et al. Cofilin-linked changes in actin filament flexibility promote severing. Biophys. J. 101, 151159 (2011).
  51. Arai, Y. et al. Tying a molecular knot with optical tweezers. Nature 399, 446448 (1999).
  52. Lenz, M., Thoresen, T., Gardel, M. L. & Dinner, A. R. Contractile units in disordered actomyosin bundles arise from F-actin buckling. Phys. Rev. Lett. 108, 238107 (2012).
  53. Murrell, M. P. & Gardel, M. L. F-actin buckling coordinates contractility and severing in a biomimetic actomyosin cortex. Proc. Natl Acad. Sci. USA 51, 2082020825 (2012).
  54. Hayakawa, K., Tatsumi, H. & Sokabe, M. Actin filaments function as a tension sensor by tension-dependent binding of cofilin to the filament. J. Cell Biol. 195, 721727 (2011).
  55. Vogel, S. K., Petrasek, Z., Heinemann, F. & Schwille, P. Myosin motors fragment and compact membrane-bound actin filaments. eLife 2, e00116 (2013).
  56. Lenz, M. Geometrical origins of contractility in disordered actomyosin networks. Phys. Rev. X 4, 041002 (2014).
  57. Thoresen, T., Lenz, M. & Gardel, M. L. Thick filament length and isoform composition determine self-organized contractile units in actomyosin bundles. Biophys. J. 104, 655665 (2013).
  58. Haviv, L., Gillo, D., Backouche, F. & Bernheim-Groswasser, A. A cytoskeletal demolition worker: myosin II acts as an actin depolymerization agent. J. Mol. Biol. 375, 325330 (2008).
  59. Pelham, R. J. & Chang, F. Actin dynamics in the contractile ring during cytokinesis in fission yeast. Nature 419, 8286 (2002).
  60. Costa, K. D., Hucker, W. J. & Yin, F. C. Buckling of actin stress fibers: a new wrinkle in the cytoskeletal tapestry. Cell. Motil. Cytoskeleton 52, 266274 (2002).
  61. Heissler, S. M. & Manstein, D. J. Nonmuscle myosin-2: mix and match. Cell. Mol. Life Sci. 70, 121 (2013).
  62. Parsons, J. T., Horwitz, A. R. & Schwartz, M. A. Cell adhesion: integrating cytoskeletal dynamics and cellular tension. Nat. Rev. Mol. Cell Biol. 11, 633643 (2010).
  63. Jordan, S. N. & Canman, J. C. Rho GTPases in animal cell cytokinesis: an occupation by the one percent. Cytoskeleton 69, 919930 (2012).
  64. Machacek, M. et al. Coordination of Rho GTPase activities during cell protrusion. Nature 461, 99103 (2009).
  65. Munro, E. & Bowerman, B. Cellular symmetry breaking during Caenorhabditis elegans development. Cold Spring Harb. Perspect. Biol. 1, a003400 (2009).
  66. Janson, L. W., Kolega, J. & Taylor, D. L. Modulation of contraction by gelation/solation in a reconstituted motile model. J. Cell Biol. 114, 10051015 (1991).
  67. Bendix, P. M. et al. A quantitative analysis of contractility in active cytoskeletal protein networks. Biophys. J. 94, 31263136 (2008).
  68. Thoresen, T., Lenz, M. & Gardel, M. L. Reconstitution of contractile actomyosin bundles. Biophys. J. 100, 26982705 (2011).
  69. Alvarado, J. et al. Molecular motors robustly drive active gels to a critically connected state. Nat. Phys. 9, 591597 (2013).
  70. Gardel, M. L. et al. Elastic behavior of cross-linked and bundled actin networks. Science 304, 13011305 (2004).
  71. Kasza, K. E. et al. Nonlinear elasticity of stiff biopolymers connected by flexible linkers. Phys. Rev. E Stat. Nonlin. Soft Matter Phys. 79, 041928 (2009).
  72. Kohler, S., Schaller, V. & Bausch, A. R. Structure formation in active networks. Nat. Mater. 10, 462468 (2011).
  73. Reymann, A.-C. et al. Nucleation geometry governs ordered actin networks structures. Nat. Mater. 9, 827832 (2010).
  74. Alexandrova, A. Y. et al. Comparative dynamics of retrograde actin flow and focal adhesions: formation of nascent adhesions triggers transition from fast to slow flow. PLoS ONE 3, e3234 (2008).
  75. Koenderink, G. H. et al. An active biopolymer network controlled by molecular motors. Proc. Natl Acad. Sci. USA 106, 1519215197 (2009).
  76. Gardel, M. L. et al. Prestressed F-actin networks cross-linked by hinged filamins replicate mechanical properties of cells. Proc. Natl Acad. Sci. USA 103, 17621767 (2006).
  77. Smith, M. A. et al. A zyxin-mediated mechanism for actin stress fiber maintenance and repair. Dev. Cell 19, 365376 (2010).
  78. Halder, G., Dupont, S. & Piccolo, S. Transduction of mechanical and cytoskeletal cues by YAP and TAZ. Nat. Rev. Mol. Cell Biol. 13, 591600 (2012).
  79. Cowan, C. R. & Hyman, A. A. Acto-myosin reorganization and PAR polarity in C. elegans. Development 134, 10351043 (2007).
  80. Liu, C. et al. Actin-mediated feedback loops in B-cell receptor signaling. Immunol. Rev. 256, 177189 (2013).
  81. Storm, C. et al. Nonlinear elasticity in biological gels. Nature 435, 191194 (2005).
  82. Gardel, M. L. et al. Stress-dependent elasticity of composite actin networks as a model for cell behavior. Phys. Rev. Lett. 96, 088102 (2006).
  83. Kasza, K. E. et al. Filamin A is essential for active cell stiffening but not passive stiffening under external force. Biophys. J. 96, 43264335 (2009).
  84. Mizuno, D., Tardin, C., Schmidt, C. F. & Mackintosh, F. C. Nonequilibrium mechanics of active cytoskeletal networks. Science 315, 370373 (2007).
  85. Pasternak, C., Spudich, J. A. & Elson, E. L. Capping of surface receptors and concomitant cortical tension are generated by conventional myosin. Nature 341, 549551 (1989).
  86. Wang, N. et al. Cell prestress. I. Stiffness and prestress are closely associated in adherent contractile cells. Am. J. Physiol. Cell Physiol. 282, C606616 (2002).
  87. Stamenovic, D., Liang, Z., Chen, J. & Wang, N. Effect of the cytoskeletal prestress on the mechanical impedance of cultured airway smooth muscle cells. J. Appl. Physiol. 92, 14431450 (2002).
  88. Balland, M., Richert, A. & Gallet, F. The dissipative contribution of myosin II in the cytoskeleton dynamics of myoblasts. Eur. Biophys. J. 34, 255261 (2005).
  89. Martens, J. C. & Radmacher, M. Softening of the actin cytoskeleton by inhibition of myosin II. Pflugers Arch. 456, 95100 (2008).
  90. Lau, A. W. et al. Microrheology, stress fluctuations, and active behavior of living cells. Phys. Rev. Lett. 91, 198101 (2003).
  91. Brangwynne, C. P. et al. Microtubules can bear enhanced compressive loads in living cells because of lateral reinforcement. J. Cell Biol. 173, 733741 (2006).
  92. Fakhri, N. et al. High-resolution mapping of intracellular fluctuations using carbon nanotubes. Science 344, 10311035 (2014).
  93. Manneville, J. B., Bassereau, P., Levy, D. & Prost, J. Activity of transmembrane proteins induces magnification of shape fluctuations of lipid membranes. Phys. Rev. Lett. 82, 43564359 (1999).
  94. Betz, T., Lenz, M., Joanny, J. F. & Sykes, C. ATP-dependent mechanics of red blood cells. Proc. Natl Acad. Sci. USA 106, 1532015325 (2009).
  95. le Duc, Q. et al. Vinculin potentiates E-cadherin mechanosensing and is recruited to actin-anchored sites within adherens junctions in a myosin II-dependent manner. J. Cell Biol. 189, 11071115 (2010).
  96. Heisenberg, C.-P. & Bellaïche, Y. Forces in tissue morphogenesis and patterning. Cell 153, 948962 (2013).
  97. Sonnemann, K. J. & Bement, W. M. Wound repair: toward understanding and integration of single-cell and multicellular wound responses. Annu. Rev. Cell Dev. Biol. 27, 237263 (2011).
  98. Friedl, P. & Gilmour, D. Collective cell migration in morphogenesis, regeneration and cancer. Nat. Rev. Mol. Cell Biol. 10, 445457 (2009).
  99. Sedzinski, J. et al. Polar actomyosin contractility destabilizes the position of the cytokinetic furrow. Nature 476, 462466 (2011).
  100. Tinevez, J.-Y. et al. Role of cortical tension in bleb growth. Proc. Natl Acad. Sci. 106, 1858118586 (2009).
  101. Rubinstein, B. et al. Actin–myosin viscoelastic flow in the keratocyte lamellipod. Biophys. J. 97, 18531863 (2009).
  102. Kruse, K., Joanny, J. F., Julicher, F. & Prost, J. Contractility and retrograde flow in lamellipodium motion. Phys. Biol. 3, 130137 (2006).
  103. Mertz, A. F. et al. Cadherin-based intercellular adhesions organize epithelial cell–matrix traction forces. Proc. Natl Acad. Sci. USA 110, 842847 (2012).
  104. Goehring, N. W. et al. Polarization of PAR proteins by advective triggering of a pattern-forming system. Science 334, 11371141 (2011).
  105. Oakes, P. W., Banerjee, S., Marchetti, M. C. & Gardel, M. L. Geometry regulates traction stresses in adherent cells. Biophys. J. 107, 825833 (2014).
  106. Guthardt Torres, P., Bischofs, I. B. & Schwarz, U. S. Contractile network models for adherent cells. Phys. Rev. E Stat. Nonlin. Soft Matter Phys. 85, 011913 (2012).
  107. Howard, J. Mechanics of Motor Proteins and the Cytoskeleton (Sinauer Associates, 2001).
  108. Yao, Norman, Y. et al. Stress-enhanced gelation: A dynamic nonlinearity of elasticity. Phys. Rev. Lett. 110, 018103 (2013).
  109. Verkhovsky, A. B., Svitkina, T. M. & Borisy, G. G. Self-polarization and directional motility of cytoplasm. Curr. Biol. 9, 1120 (1999).
  110. Sun, S. X., Walcott, S. & Wolgemuth, C. W. Cytoskeletal cross-linking and bundling in motor-independent contraction. Curr. Biol. 20, R649R654 (2010).
  111. Ramaswamy, S. The mechanics and statistics of active matter. Annu. Rev. Condensed Matter Phys. 1, 323345 (2010).
  112. Bartles, J. R. Parallel actin bundles and their multiple actin-bundling proteins. Curr. Opin. Cell Biol. 12, 7278 (2000).
  113. Kohler, S. & Bausch, A. R. Contraction mechanisms in composite active actin networks. PLoS ONE 7, e39869 (2012).
  114. Kane, R. E. Interconversion of structural and contractile actin gels by insertion of myosin during assembly. J. Cell Biol. 97, 17451752 (1983).
  115. Backouche, F., Haviv, L., Groswasser, D. & Bernheim-Groswasser, A. Active gels: dynamics of patterning and self-organization. Phys. Biol. 3, 264273 (2006).
  116. Aratyn, Y. S., Schaus, T. E., Taylor, E. W. & Borisy, G. G. Intrinsic dynamic behavior of fascin in filopodia. Mol. Biol. Cell 18, 39283940 (2007).
  117. Wang, K., Ash, J. F. & Singer, S. J. Filamin, a new high-molecular-weight protein found in smooth muscle and non-muscle cells. Proc. Natl Acad. Sci. USA 72, 44834486 (1975).
  118. Biro, Maté et al. Cell cortex composition and homeostasis resolved by integrating proteomics and quantitative imaging. Cytoskeleton 70, 741754 (2013).
  119. Schmoller, K. M., Lieleg, O. & Bausch, A. R. Structural and viscoelastic properties of actin/filamin networks: cross-linked versus bundled networks. Biophys. J. 97, 8389 (2009).
  120. Kasza, K. E. et al. Actin filament length tunes elasticity of flexibly cross-linked actin networks. Biophys. J. 99, 10911100 (2010).
  121. Kohler, S., Schmoller, K. M., Crevenna, A. H. & Bausch, A. R. Regulating contractility of the actomyosin cytoskeleton by pH. Cell Rep. 2, 433439 (2012).
  122. Goldmann, W. H. & Isenberg, G. Analysis of filamin and α-actinin binding to actin by the stopped flow method. FEBS Lett. 336, 408410 (1993).
  123. Ebashi, S. & Ebashi, F. α-actinin, a new structural protein from striated muscle. I. Preparation and action on actomyosin-ATP interaction. J. Biochem. 58, 712 (1965).
  124. Edlund, M., Lotano, M. A. & Otey, C. A. Dynamics of α-actinin in focal adhesions and stress fibers visualized with α-actinin–green fluorescent protein. Cell. Motil. Cytoskeleton 48, 190200 (2001).
  125. Sanger, J. M., Mittal, B., Pochapin, M. B. & Sanger, J. W. Stress fiber and cleavage furrow formation in living cells microinjected with fluorescently labeled α-actinin. Cell. Motil. Cytoskeleton 7, 209220 (1987).
  126. Falzone, T. T., Lenz, M., Kovar, D. R. & Gardel, M. L. Assembly kinetics determine the architecture of α-actinin crosslinked F-actin networks. Nat. Commun. 3, 861 (2012).
  127. Field, C. M. & Alberts, B. M. Anillin, a contractile ring protein that cycles from the nucleus to the cell cortex. J. Cell Biol. 131, 165178 (1995).
  128. Schaller, V. et al. Crosslinking proteins modulate the self-organization of driven systems. Soft Matter 9, 72297233 (2013).
  129. Kinoshita, M. et al. Self- and actin-templated assembly of Mammalian septins. Dev. Cell 3, 791802 (2002).
  130. Reichl, E. M. et al. Interactions between myosin and actin crosslinkers control cytokinesis contractility dynamics and mechanics. Curr. Biol. 18, 471480 (2008).
  131. Weber, I. et al. Two-step positioning of a cleavage furrow by cortexillin and myosin II. Curr. Biol. 10, 501506 (2000).
  132. Yin, H. L. & Stossel, T. P. Control of cytoplasmic actin gel–sol transformation by gelsolin, a calcium-dependent regulatory protein. Nature 281, 583586 (1979).
  133. Murrell, M. et al. Spreading dynamics of biomimetic actin cortices. Biophys. J. 100, 14001409 (2011).
  134. Murrell, M. & Gardel, M. L. Actomyosin sliding is attenuated in contractile biomimetic cortices. Mol. Biol. Cell 25, 18451853 (2014).
  135. Carvalho, K. et al. Cell-sized liposomes reveal how actomyosin cortical tension drives shape change. Proc. Natl Acad. Sci. USA 110, 1645616461 (2013).

Download references

Author information

Affiliations

  1. Department of Biomedical Engineering, Yale University, New Haven, Connecticut 06520, USA.

    • Michael Murrell
  2. Systems Biology Institute, Yale University, West Haven, Connecticut 06516, USA.

    • Michael Murrell
  3. Department of Physics, Institute for Biophysical Dynamics and James Franck Institute, University of Chicago, Chicago, Illinois 60637, USA.

    • Patrick W. Oakes &
    • Margaret L. Gardel
  4. Université Paris-Sud, CNRS, LPTMS, UMR 8626, Orsay 91405, France.

    • Martin Lenz

Competing interests statement

The authors declare no competing interests.

Corresponding author

Correspondence to:

Author details

  • Michael Murrell

    Michael Murrell obtained his Ph.D. from Masachussetts Institute for Technology, Cambridge, USA, and is currently an assistant professor at the Systems Biology Institute and Biomedical Engineering Department at Yale University, New Haven Connecticut, USA. His interests lie in understanding the mechanical principles that drive major cellular life processes through the design and engineering of novel biomimetic systems.

  • Patrick W. Oakes

    Patrick W. Oakes obtained his Ph.D. from Brown University, Providence, Rhode Island, USA, and is currently a postdoctoral scholar in the laboratory of Margaret Gardel at the University of Chicago, Illinois, USA. His studies are focused on how cells regulate their contractile behavior and how they interact with their extracellular environment.

  • Martin Lenz

    Martin Lenz is a researcher in the Theoretical Physics division of the Centre National de la Recherche Scientifique (CNRS) in France. His group, located in Orsay, studies the relationships between protein-scale processes and resulting cell- or tissue-scale biomechanical properties using mathematical and computational tools from soft matter and statistical physics.

  • Margaret L. Gardel

    Margaret L. Gardel obtained her Ph.D. from Harvard University, Cambridge, Massachussetts, USA, and is currently an associate professor at University of Chicago, Illinois, USA, in the Physics Department, James Franck Institute and Institute for Biophysical Dyanmics. Her laboratory studies the biophysical regulation of cell adhesion, motion and shape by cytoskeletal assemblies.

Additional data