Actin Filament


What are actin filaments[Edit]

Actin filaments (F-actin) are linear polymers of globular actin (G-actin) subunits and occur as microfilaments in the cytoskeleton and as thin filaments, which are part of the contractile apparatus, in muscle and nonmuscle cells (see contractile bundles). They commonly underlie the plasma membrane and are typically assembled at the cell periphery from adhesion sites or sites of membrane extension. Actin filaments can create a number of linear bundles, two-dimensional networks, and three-dimensional gels, and actin binding proteins can influence the specific structure the filaments will form. 

Although a large body of work suggests that F-actin can exist in multiple states, in general, an actin filament has a total rise of 27.3 Å between subunits on adjacent strands and a rotation of 166.15° around the axis [1]. An actin filament is flexible, has a helical repeat every 37 nm, ranges from 5-9 nm in diameter, and has 13 actin subunits between each cross-over point (produced by the ‘crossing over’ of the two long-pitch actin helices) (reviewed in [2, 3]). An exception is the case of sperm, where a crosslinker protein, scruin, maintains the actin in a coiled state and extends upon activation. In the acrosomal actin rotation angle of one of the strands is 167.15° while the other strand is tilted more than usual to fit into an asymmetric helix [4].

Actin Filament Polarity[Edit]

The polarity of an actin filament is visualized by the binding of the myosin subfragment (S1) to the filament, which creates barbed (+) and pointed (-) ends on the filament [5]. 
Figure 1. Structure of an actin filament showing the barbed (or plus) and pointed (or minus) ends: This diagram illustrates the molecular organization of actin and provides examples for how an actin filament is represented in figures throughout this resource. Early models for actin filaments were constructed by fitting the filament x-ray crystal structure to the atomic structure of actin monomers [6](reviewed in [2]) while more recent models use a number of different approaches [7].
When all actin subunits are bound by myosin S1, the filament appears coated with arrowheads that all point towards one end of the filament [8, 9]. The barbed end structure with bound capping protein has been determined [10] and polarity ascertained by novel single-particle image analysis methods applied to electromicrographs of in vitro protein preparations [11]. Polarity has since also been determined from cryo-electron tomograms of vitreously frozen cells [12]. The (+) end of filaments generally have a high concentration of F-actin-ATP and denotes the growing end of an actin filament.

The (-) end generally has a high concentration of F-actin-ADP and denotes the shrinking end of an actin filament. Most actin filaments are arranged with the barbed end toward the cellular membrane and the pointed end toward the cellular interior. The length of actin filaments can be adjusted by the activity of actin binding proteins and by altering the rate of actin treadmilling.

Distribution of actin filaments in cells and tissues[Edit]

Actin filaments are widely distributed throughout cells, forming a range of cytoskeletal structures and contributing to an even broader range of processes. Some of the functions are widely observed across many cell types while in other cases actin filaments may contribute to processes that are cell type specific.

1. In polarized cells and tissues:

Figure 2. Actin filament distribution in cells and tissues
Actin filaments are crucial for tissue organization and for establishing cell polarity and cohesion among epithelial cells. For example, a core of actin filaments provides microvilli structural support and enables them to increase their surface area and nutrient-absorbing capacity. These structures are found on the apical surface of epithlial cells lining the small intestine. In another example, the integrity of epithelial cell layers or sheets is maintained by a belt of actin filaments (i.e. adhesion belt). This belt links the cytoskeleton of adjacent cells. Also, certain cells use actin filament rigidity to sense vibrations, such as those found bundled on the surface of hair cells in the inner ear (called stereocilia, not shown), which tilt in response to sound. Although the actin bundles in stereocilia are stable for the lifetime of a cell (which can be decades), the individual actin filaments are continuously remodeled and replaced once every 48 hours (on average).

2. In motility-related structures:

Figure 3. Cell cortex (aka cortical actin, actin cortex): (A) Cortical actin filaments (shown in red) are concentrated just beneath the plasma membrane in most cell types. (B) Migrating fibroblasts grown in 2D tissue culture have more cortical filaments on the dorsal (upper) surface than the ventral (lower) surface and they are concentrated towards the trailing edges [13]
Actin filaments are the primary cytoskeletal component to drive cell motility. Here, actin filaments found in membrane protrusions such as filopodia and lamellipodia rapidly assemble and disassemble. These cellular structures are essential in cell migration and are predominately found at the leading edge of a moving cell. They also allow the cell to probe or sense its microenvironment. Actin filaments are also found at the trailing edge in the form of the cell cortex, which lies adjacent to the plasma membrane. More stable arrays of actin filaments, such as those found in stress fibers, allow a cell to brace against the underlying surface. Actin-associated myosin motor proteins use ATP hydrolysis to exert forces against the stress fibers during muscle contraction; the energy of hydrolysis can also be converted to tensile forces at the trailing cell edge to promote edge retraction in motile cells.

3. During cell division:

Actin-based motile structures are disassembled before cell division, which causes the cell to stop moving and become more rounded. More stable actin bundles remain polarized and contribute to the orientation of the microtubule network that serves as the mitotic spindle. The proper assembly, orientation, and contraction of an actin filament ring (i.e. contractile ring) serves to pinch and separate the daughter cells during cell division.

4. During reproduction:

Actin filaments are an important structural feature in the head of sperm. The rapid relaxing of coiled actin filaments during the acrosome reaction [4] allows the sperm head to penetrate further into the egg. Microtubules influence actin assembly to help organize the polarized actin network. 

Actin Filament Function[Edit]

Several biological processes related to cell shape and movement depend on actin filaments (reviewed in [14]). Some keys functions are:

* To form the dynamic cytoskeleton, which gives structural support to cells and links the interior of the cell with its surroundings. Forces acting on the actin cytoskeleton are translated and transmitted by signaling pathways to convey information about the external environment.
* To allow cell motility. For example, through the formation and function of Filopodia or Lamellipodia.
* During mitosis, intracellular organelles are transported by motor proteins to the daughter cells along actin cables
* In muscle cells, actin filaments are aligned and myosin proteins generate forces on the filaments to support muscle contraction. These complexes are known as ‘thin filaments’. 
* In non-muscle cells, actin filaments form a track system for cargo transport that is powered by non-conventional myosins such as myosin V and VI. Non-conventional myosins use the energy from ATP hydrolysis to transport cargo (such as vesicles and organelles) at rates much faster than diffusion.

Thin filament

In certain cases, actin filaments are assembled with, and stabilized by, accessory proteins into higher order contractile structures such as stress fibers (nonmuscle cells) or contractile bundles (muscle cells). The dynamic association of tropomyosin and troponin with actin filaments stabilizes the actin filament (collectively termed “thin filament”) to be functional in various contexts.

Figure 4. Tropomyosin stabilizes thin filaments: TM binds to the side of adjacent actin subunits along the groove of the helix to stabilize and stiffen the actin filament [15]. TM also prevents other proteins from accessing the filament; this inhibition is essential for regulating muscle contraction [16]. TN controls the positioning of TM along the groove of the actin filament.
A single tropomyosin binds to the side of adjacent actin subunits and extends over approximately seven actin monomers [15]. End-to-end binding of tropomyosins produces a continuous strand of tropomyosin polymers along the groove of the actin helix which allows their cooperative movement [17]. Tropomyosin isoforms stabilize actin filaments [18] and occupy the same binding sites on actin that are used by known regulators of actin filaments (e.g. ADF/cofilin [19]) (reviewed in [20, 21]). 

Focus on contractile machinery

Troponin, a three-peptide complex, is thought to trap tropomyosin in a calcium-dependent fashion at a position that inhibits myosin bundles from accessing the actin filaments; calcium binding to troponin allows a conformational restructuring of tropomyosin that leaves the myosin-binding sites on the thin filaments exposed [22, 23, 24, 25]. Subsequent binding of the myosin thick filaments augments movement of tropomyosin away from the actin filament and full exposure of the myosin binding sites [24]. However, control of tropomyosin-binding to myosin thick filaments may be independent of troponin presence [18]; smooth muscle cells and many non-muscle cells lack troponin.

Thus TM regulates both the association of myosin bundles with actin filaments [25] and their ATPase kinetics (reviewed in[26]). It is likely that TM isoforms from different tissues or cell types may have specific effects on actomyosin ATPase activity and cytoskeletal functions [18, 27]

Actin filament polymerization generates protrusive force[Edit]

Figure 5. Actin polymerization produces force for movement: Pathogenic intracellular bacteria use mainly host-derived components to facilitate movement through (and between) cells. The bacterial ActA protein on the surface of the bacterium (e.g. Listeria monocytogenes) recruits and activates the host components needed for actin polymerization (e.g. Ena/VASP, ATP-actin). The actin filaments are assembled at the plus end nearest the bacteria membrane and the forces generated by filament assembly are translated into bacterium movement. The resulting actin comet tail is likely comprised of branched arrays of actin filaments as seen from simulation of experimental results [28, 29, 30]. The dynamics of filament assembly and turnover (and therefore bacterium motility) are controlled by capping protein, profilin, ADF/Cofilin and Arp2/3 complex.
It has been known for some time that actin polymerization produces most of the driving force for membrane protrusion [31, 32, 33] (reviewed in [34, 35, 36, 37]). Experiments to identify the mechanism and minimum components for actin-based protrusion have used both pathogenic intracellular bacteria in in vitro reconstitution experiments [33] and baculovirus in host [38] (see Figure below).

In cells, actin filaments are initiated with their barbed ends oriented towards the plasma membrane, with ATP hydrolysis facilitating filament growth. Polymerization is favored towards the cell front and disassembly occurs more frequently at the rear (reviewed in [39]). However, only a small fraction of the overall free energy of nucleotide hydrolysis is needed to modulate G-actin monomer binding. The remaining energy is translated into a protrusive force that deforms the plasma membrane in a particular direction [40, 41, 42, 43] (reviewed in [35]).

The propulsive network is self-organizing and filaments with a particular orientation,with respect to the membrane, will assemble at the maximal velocity and be preferentially chosen for elongation [44]. Similarly, cell shape and migration speed is determined by a dynamic steady state that is self-organizing [45]. As actin filaments grow, they remain fixed within the cytoskeletal network (reviewed in [46]).

Factors that regulate the protrusive force of actin filaments:

Factors that influence the concentration of free G-actin (e.g. profilin [47]) or ATP-binding and hydrolysis on actin (reviewed in [38]) will promote filament assembly and membrane protrusion.

The microtubule and intermediate filament networks play a key role in regulating the global deposition pattern of the actin filaments, therefore, they will also influence membrane protrusion dynamics [48] (reviewed in [44, 45, 49, 50, 51]).

Membrane tension- In order for a cell to extend its leading edge forward, the cell must overcome resisting forces. Motile cells in living systems experience external forces from the surrounding material (usually ECM), while the major force resisting extension for cells in tissue culture are tensile forces within the plasma membrane. Biophysical models [44, 52, 53] (reviewed in [54, 55]) and experiments with live cells have shown that the membrane extension rate is directly dependent on the membrane tension: elevated tension lowers cell membrane extension and motility, regardless of whether the tension is applied externally (e.g. stretching) or internally (e.g. contraction of stress fibers) [43, 56, 57].

Actin filaments are a force-sensing conduit for both internal and external forces[Edit]

Internal forces 

The orientation of individual actin filaments in the cytoskeleton is a force-driven evolutionary process [58] that contributes to the elastic behavior of the network and influences whether a filament will deform by compression, bending or extension [33, 59].Cross-linked actin networks initially become more elastic under low force as a result of filament resistance to the direction of compression [60]. As the force increases, individual filaments inherently resist being compressed and/or cross-linking proteins become more extended, which causes the cytoskeleton network to become more rigid [61, 62]; cell stiffening has also been correlated with actin recruitment [63]. Each of these processes in turn, can influence mechanosensors that are linked to the actin network to induce subsequent mechanotransduction events. Although forces ~ 400 pN along the filament axis will break actin filaments, much higher forces are generated in migrating cells (reviewed in[64]); this raises the question “how does a cell alleviate the forces on the internal actin network?” The answer lies in the proteins that are linked to the actin: extreme stress on the filaments causes buckling and reduced elasticity of the network in a process known as ‘stress softening’ [65]. Softening is attributed to either filament fracturing, which leads to new uncapped ends for filament assembly/disassembly, or to softening, which may be due to unbinding of flexible protein crosslinkers [60, 66]. In addition, because the filaments are interconnected to the rest of the cytoskeleton, buckled filaments don’t collapse and the process can be reversed when the stress is removed [67] (reviewed in [68, 69]).

External or applied forces 

The actin cytoskeleton is physically connected with the cell exterior, e.g., ECM or other cells, through a multiprotein complex known as the focal adhesion complex. Interactions between cell surface molecules, e.g., integrins, and the actin cytoskeleton are bidirectional, with the focal adhesion complex forming the link between them [70]. Actin filaments and their associated focal adhesion complexes act as information handling machines or mechanosensors: they convert both the strength of the adhesion and the tensile forces along the linked network of actin filaments (and associated proteins) into biochemical signals that control actin extension and cell migration (reviewed in [69, 71, 72,73, 74, 75]). Focal adhesions are subject to continuous pulling forces [76, 77] and the force differences due to external stress [78] or the chemical nature (reviewed in [79]), rigidity [80,81], and topography of the exterior components [82, 83, 84] (reviewed in [46, 85, 86]) will influence the assembly and organization of the actin cytoskeleton [84, 45, 87, 88, 89].

Actin filaments form higher-order assemblies that produce and respond to force[Edit]

Many different types of cells use actin filaments to generate tensional forces and to exert traction forces on their adhesions and linked ECM (or other cells); these processes cause resting tensile force (reviewed in [68, 90]). Local stress or force differences that occur both internally and externally are transduced through the actin filaments and focused onto mechanosensors throughout the interlinked system rather than being limited to local deformation (reviewed in [91]); this process leads to mechanotransduction events that influence the cell shape and/or motility (reviewed in [92]). 

Cells exert traction forces on the ECM and generate tension at focal adhesions through actin stress fibers, which are higher-order structures in the cytoplasm that consist of parallel contractile bundles of actin and myosin filaments. Stress fibers are linked at their ends to the ECM through focal adhesion complexes. Cell tension is generated along the actin filaments by the movement of myosin II motor proteins along the filaments (see contractile bundles). The elasticity of the actin network appears to be inherent to actin filaments and is independent of myosin II motor activity [65]. Forces produced by the contraction of stress fibers not only helps the cell body to translocate during migration [84, 93], but they also serve as a vital “inside-out” feedback system to regulate actin filament initiation [94], cell growth and motility [95, 96], and formation/maturation of focal adhesion complexes [97, 98]. Forces produced by stress fibers also stabilize the cell structure and contribute to establishing the cell polarity [93] and they help determine the cell fate [76, 99]. There is no evidence that forces influence stress fiber contractile activity by increasing the exchange of ADP for ATP on myosin, however, force can weakly increase the release of the myosin heads from the actin filaments (reviewed in [64]).

References

  1. Egelman EH., Francis N., DeRosier DJ. F-actin is a helix with a random variable twist. Nature 1982; 298(5870). [PMID: 7201078]
  2.  The structure of F-actin. J. Muscle Res. Cell. Motil. 1985; 6(2). [PMID: 3897278]
  3.  A tale of two polymers: new insights into helical filaments. Nat. Rev. Mol. Cell Biol. 2003; 4(8). [PMID: 12923524]
  4. Schmid MF., Sherman MB., Matsudaira P., Chiu W. Structure of the acrosomal bundle. Nature 2004; 431(7004). [PMID: 15343340]
  5. Moore PB., Huxley HE., DeRosier DJ. Three-dimensional reconstruction of F-actin, thin filaments and decorated thin filaments. J. Mol. Biol. 1970; 50(2). [PMID: 5476917]
  6. Holmes KC., Popp D., Gebhard W., Kabsch W. Atomic model of the actin filament. Nature 1990; 347(6288). [PMID: 2395461]
  7. Oda T., Stegmann H., Schröder RR., Namba K., Maéda Y. Modeling of the F-actin structure. Adv. Exp. Med. Biol. 2007; 592. [PMID: 17278381]
  8. Mooseker MS., Tilney LG. Organization of an actin filament-membrane complex. Filament polarity and membrane attachment in the microvilli of intestinal epithelial cells. J. Cell Biol. 1975; 67(3). [PMID: 1202021]
  9. Begg DA., Rodewald R., Rebhun LI. The visualization of actin filament polarity in thin sections. Evidence for the uniform polarity of membrane-associated filaments. J. Cell Biol. 1978; 79(3). [PMID: 569662]
  10. Narita A., Takeda S., Yamashita A., Maéda Y. Structural basis of actin filament capping at the barbed-end: a cryo-electron microscopy study. EMBO J. 2006; 25(23). [PMID: 17110933]
  11. Narita A., Maéda Y. Molecular determination by electron microscopy of the actin filament end structure. J. Mol. Biol. 2007; 365(2). [PMID: 17059832]
  12. Urban E., Jacob S., Nemethova M., Resch GP., Small JV. Electron tomography reveals unbranched networks of actin filaments in lamellipodia. Nat. Cell Biol. 2010; 12(5). [PMID: 20418872]
  13. Makar AB., McMartin KE., Palese M., Tephly TR. Formate assay in body fluids: application in methanol poisoning. Biochem Med 1975; 13(2). [PMID: 1]
  14. Pollard TD., Cooper JA. Actin, a central player in cell shape and movement. Science 2009; 326(5957). [PMID: 19965462]
  15. Ebashi S., Endo M. Calcium ion and muscle contraction. Prog. Biophys. Mol. Biol. 1968; 18. [PMID: 4894870]
  16. Huckaba TM., Lipkin T., Pon LA. Roles of type II myosin and a tropomyosin isoform in retrograde actin flow in budding yeast. J. Cell Biol. 2006; 175(6). [PMID: 17178912]
  17. Flicker PF., Phillips GN., Cohen C. Troponin and its interactions with tropomyosin. An electron microscope study. J. Mol. Biol. 1982; 162(2). [PMID: 7161805]
  18. Lehman W., Hatch V., Korman V., Rosol M., Thomas L., Maytum R., Geeves MA., Van Eyk JE., Tobacman LS., Craig R. Tropomyosin and actin isoforms modulate the localization of tropomyosin strands on actin filaments. J. Mol. Biol. 2000; 302(3). [PMID: 10986121]
  19. Ono S., Ono K. Tropomyosin inhibits ADF/cofilin-dependent actin filament dynamics. J. Cell Biol. 2002; 156(6). [PMID: 11901171]
  20.  F-actin-binding proteins. Curr. Opin. Struct. Biol. 1998; 8(2). [PMID: 9631289]
  21. Bryce NS., Schevzov G., Ferguson V., Percival JM., Lin JJ., Matsumura F., Bamburg JR., Jeffrey PL., Hardeman EC., Gunning P., Weinberger RP. Specification of actin filament function and molecular composition by tropomyosin isoforms. Mol. Biol. Cell 2003; 14(3). [PMID: 12631719]
  22. Lehman W., Craig R., Vibert P. Ca(2+)-induced tropomyosin movement in Limulus thin filaments revealed by three-dimensional reconstruction. Nature 1994; 368(6466). [PMID: 8107884]
  23. Lehman W., Vibert P., Uman P., Craig R. Steric-blocking by tropomyosin visualized in relaxed vertebrate muscle thin filaments. J. Mol. Biol. 1995; 251(2). [PMID: 7643394]
  24. Vibert P., Craig R., Lehman W. Steric-model for activation of muscle thin filaments. J. Mol. Biol. 1997; 266(1). [PMID: 9054965]
  25. Xu C., Craig R., Tobacman L., Horowitz R., Lehman W. Tropomyosin positions in regulated thin filaments revealed by cryoelectron microscopy. Biophys. J. 1999; 77(2). [PMID: 10423443]
  26.  Tropomyosins as discriminators of myosin function. Adv. Exp. Med. Biol. 2008; 644. [PMID: 19209828]
  27. Lehman W., Szent-Györgyi G. Activation of the adenosine triphosphatase of Limulus polyphemus actomyosin by tropomyosin. J. Gen. Physiol. 1972; 59(4). [PMID: 4260494]
  28. Smith RJ., Bryant RG. Metal substitutions incarbonic anhydrase: a halide ion probe study. Biochem. Biophys. Res. Commun. 1975; 66(4). [PMID: 3]
  29. Moroi K., Sato T. Comparison between procaine and isocarboxazid metabolism in vitro by a liver microsomal amidase-esterase. Biochem. Pharmacol. 1975; 24(16). [PMID: 8]
  30. Thornton JA., Harrison MJ. Letter: Duration of action of AH8165. Br J Anaesth 1975; 47(9). [PMID: 28]
  31.  Microfilament or microtubule assembly or disassembly against a force. Proc. Natl. Acad. Sci. U.S.A. 1981; 78(9). [PMID: 6946498]
  32. Peskin CS., Odell GM., Oster GF. Cellular motions and thermal fluctuations: the Brownian ratchet. Biophys. J. 1993; 65(1). [PMID: 8369439]
  33. Loisel TP., Boujemaa R., Pantaloni D., Carlier MF. Reconstitution of actin-based motility of Listeria and Shigella using pure proteins. Nature 1999; 401(6753). [PMID: 10524632]
  34. Borisy GG., Svitkina TM. Actin machinery: pushing the envelope. Curr. Opin. Cell Biol. 2000; 12(1). [PMID: 10679366]
  35.  The polymerization motor. Traffic 2000; 1(1). [PMID: 11208055]
  36. Pollard TD., Borisy GG. Cellular motility driven by assembly and disassembly of actin filaments. Cell 2003; 112(4). [PMID: 12600310]
  37. Le Clainche C., Carlier MF. Regulation of actin assembly associated with protrusion and adhesion in cell migration. Physiol. Rev. 2008; 88(2). [PMID: 18391171]
  38. Ohkawa T., Volkman LE., Welch MD. Actin-based motility drives baculovirus transit to the nucleus and cell surface. J. Cell Biol. 2010; 190(2). [PMID: 20660627]
  39. Carlier MF., Pantaloni D. Control of actin dynamics in cell motility. J. Mol. Biol. 1997; 269(4). [PMID: 9217250]
  40. Oliver T., Dembo M., Jacobson K. Separation of propulsive and adhesive traction stresses in locomoting keratocytes. J. Cell Biol. 1999; 145(3). [PMID: 10225959]
  41. Elson EL., Felder SF., Jay PY., Kolodney MS., Pasternak C. Forces in cell locomotion. Biochem. Soc. Symp. 1999; 65. [PMID: 10320946]
  42. Dickinson RB., Caro L., Purich DL. Force generation by cytoskeletal filament end-tracking proteins. Biophys. J. 2004; 87(4). [PMID: 15454475]
  43. Marcy Y., Prost J., Carlier MF., Sykes C. Forces generated during actin-based propulsion: a direct measurement by micromanipulation. Proc. Natl. Acad. Sci. U.S.A. 2004; 101(16). [PMID: 15079054]
  44. Mogilner A., Oster G. Cell motility driven by actin polymerization. Biophys. J. 1996; 71(6). [PMID: 8968574]
  45. Keren K., Pincus Z., Allen GM., Barnhart EL., Marriott G., Mogilner A., Theriot JA. Mechanism of shape determination in motile cells. Nature 2008; 453(7194). [PMID: 18497816]
  46. Vogel V., Sheetz M. Local force and geometry sensing regulate cell functions. Nat. Rev. Mol. Cell Biol. 2006; 7(4). [PMID: 16607289]
  47. Kang F., Purich DL., Southwick FS. Profilin promotes barbed-end actin filament assembly without lowering the critical concentration. J. Biol. Chem. 1999; 274(52). [PMID: 10601251]
  48. Pan Y., Jing R., Pitre A., Williams BJ., Skalli O. Intermediate filament protein synemin contributes to the migratory properties of astrocytoma cells by influencing the dynamics of the actin cytoskeleton. FASEB J. 2008; 22(9). [PMID: 18509200]
  49. Fuchs E., Yang Y. Crossroads on cytoskeletal highways. Cell 1999; 98(5). [PMID: 10490093]
  50. Goode BL., Drubin DG., Barnes G. Functional cooperation between the microtubule and actin cytoskeletons. Curr. Opin. Cell Biol. 2000; 12(1). [PMID: 10679357]
  51.  Actin and microtubules in cell motility: which one is in control? Traffic 2004; 5(7). [PMID: 15180824]
  52.  Biophysics of the leading lamella. Cell Motil. Cytoskeleton 1988; 10(1-2). [PMID: 3052865]
  53. Mogilner A., Oster G. Force generation by actin polymerization II: the elastic ratchet and tethered filaments. Biophys. J. 2003; 84(3). [PMID: 12609863]
  54.  On the edge: modeling protrusion. Curr. Opin. Cell Biol. 2006; 18(1). [PMID: 16318917]
  55. Rangarajan R., Zaman MH. Modeling cell migration in 3D: Status and challenges. Cell Adh Migr undefined; 2(2). [PMID: 19262098]
  56. Sheetz MP., Dai J. Modulation of membrane dynamics and cell motility by membrane tension. Trends Cell Biol. 1996; 6(3). [PMID: 15157483]
  57. Karl I., Bereiter-Hahn J. Tension modulates cell surface motility: A scanning acoustic microscopy study. Cell Motil. Cytoskeleton 1999; 43(4). [PMID: 10423275]
  58. Maly IV., Borisy GG. Self-organization of a propulsive actin network as an evolutionary process. Proc. Natl. Acad. Sci. U.S.A. 2001; 98(20). [PMID: 11572984]
  59. Kroy K., Frey E. Force-Extension Relation and Plateau Modulus for Wormlike Chains. Phys. Rev. Lett. 1996; 77(2). [PMID: 10062418]
  60. Gardel ML., Shin JH., MacKintosh FC., Mahadevan L., Matsudaira P., Weitz DA. Elastic behavior of cross-linked and bundled actin networks. Science 2004; 304(5675). [PMID: 15166374]
  61. Footer MJ., Kerssemakers JW., Theriot JA., Dogterom M. Direct measurement of force generation by actin filament polymerization using an optical trap. Proc. Natl. Acad. Sci. U.S.A. 2007; 104(7). [PMID: 17277076]
  62. Johnson CP., Tang HY., Carag C., Speicher DW., Discher DE. Forced unfolding of proteins within cells. Science 2007; 317(5838). [PMID: 17673662]
  63. Icard-Arcizet D., Cardoso O., Richert A., Hénon S. Cell stiffening in response to external stress is correlated to actin recruitment. Biophys. J. 2008; 94(7). [PMID: 18178644]
  64. Khan S., Sheetz MP. Force effects on biochemical kinetics. Annu. Rev. Biochem. 1997; 66. [PMID: 9242924]
  65. Chaudhuri O., Parekh SH., Fletcher DA. Reversible stress softening of actin networks. Nature 2007; 445(7125). [PMID: 17230186]
  66. Gardel ML., Nakamura F., Hartwig JH., Crocker JC., Stossel TP., Weitz DA. Prestressed F-actin networks cross-linked by hinged filamins replicate mechanical properties of cells. Proc. Natl. Acad. Sci. U.S.A. 2006; 103(6). [PMID: 16446458]
  67. Ingber DE., Folkman J. Mechanochemical switching between growth and differentiation during fibroblast growth factor-stimulated angiogenesis in vitro: role of extracellular matrix. J. Cell Biol. 1989; 109(1). [PMID: 2473081]
  68.  Tensegrity-based mechanosensing from macro to micro. Prog. Biophys. Mol. Biol. undefined; 97(2-3). [PMID: 18406455]
  69.  Cellular mechanotransduction: putting all the pieces together again. FASEB J. 2006; 20(7). [PMID: 16675838]
  70.  Bidirectional control of the inner dynamics of focal adhesions promotes cell migration. Cell Adh Migr undefined; 3(2). [PMID: 19398887]
  71. Bershadsky A., Kozlov M., Geiger B. Adhesion-mediated mechanosensitivity: a time to experiment, and a time to theorize. Curr. Opin. Cell Biol. 2006; 18(5). [PMID: 16930976]
  72. Bershadsky AD., Ballestrem C., Carramusa L., Zilberman Y., Gilquin B., Khochbin S., Alexandrova AY., Verkhovsky AB., Shemesh T., Kozlov MM. Assembly and mechanosensory function of focal adhesions: experiments and models. Eur. J. Cell Biol. 2006; 85(3-4). [PMID: 16360240]
  73.  Mechanotransduction – a field pulling together? J. Cell. Sci. 2008; 121(Pt 20). [PMID: 18843115]
  74. Lock JG., Wehrle-Haller B., Strömblad S. Cell-matrix adhesion complexes: master control machinery of cell migration. Semin. Cancer Biol. 2008; 18(1). [PMID: 18023204]
  75. Geiger B., Spatz JP., Bershadsky AD. Environmental sensing through focal adhesions. Nat. Rev. Mol. Cell Biol. 2009; 10(1). [PMID: 19197329]
  76. Griffin MA., Sen S., Sweeney HL., Discher DE. Adhesion-contractile balance in myocyte differentiation. J. Cell. Sci. 2004; 117(Pt 24). [PMID: 15522893]
  77. Gibson MC., Perrimon N. Extrusion and death of DPP/BMP-compromised epithelial cells in the developing Drosophila wing. Science 2005; 307(5716). [PMID: 15774762]
  78. Helmke BP., Davies PF. The cytoskeleton under external fluid mechanical forces: hemodynamic forces acting on the endothelium. Ann Biomed Eng 2002; 30(3). [PMID: 12051614]
  79. Hersel U., Dahmen C., Kessler H. RGD modified polymers: biomaterials for stimulated cell adhesion and beyond. Biomaterials 2003; 24(24). [PMID: 12922151]
  80. Discher DE., Janmey P., Wang YL. Tissue cells feel and respond to the stiffness of their substrate. Science 2005; 310(5751). [PMID: 16293750]
  81. Engler AJ., Sen S., Sweeney HL., Discher DE. Matrix elasticity directs stem cell lineage specification. Cell 2006; 126(4). [PMID: 16923388]
  82. Curtis A., Wilkinson C. New depths in cell behaviour: reactions of cells to nanotopography. Biochem. Soc. Symp. 1999; 65. [PMID: 10320930]
  83. Dalby MJ., Riehle MO., Johnstone H., Affrossman S., Curtis AS. In vitro reaction of endothelial cells to polymer demixed nanotopography. Biomaterials 2002; 23(14). [PMID: 12069336]
  84. Parker KK., Brock AL., Brangwynne C., Mannix RJ., Wang N., Ostuni E., Geisse NA., Adams JC., Whitesides GM., Ingber DE. Directional control of lamellipodia extension by constraining cell shape and orienting cell tractional forces. FASEB J. 2002; 16(10). [PMID: 12153987]
  85. Curtis A., Riehle M. Tissue engineering: the biophysical background. Phys Med Biol 2001; 46(4). [PMID: 11324976]
  86. Spatz JP., Geiger B. Molecular engineering of cellular environments: cell adhesion to nano-digital surfaces. Methods Cell Biol. 2007; 83. [PMID: 17613306]
  87. Singhvi R., Kumar A., Lopez GP., Stephanopoulos GN., Wang DI., Whitesides GM., Ingber DE. Engineering cell shape and function. Science 1994; 264(5159). [PMID: 8171320]
  88. Chen CS., Mrksich M., Huang S., Whitesides GM., Ingber DE. Geometric control of cell life and death. Science 1997; 276(5317). [PMID: 9162012]
  89. Parekh SH., Chaudhuri O., Theriot JA., Fletcher DA. Loading history determines the velocity of actin-network growth. Nat. Cell Biol. 2005; 7(12). [PMID: 16299496]
  90.  Tensegrity: the architectural basis of cellular mechanotransduction. Annu. Rev. Physiol. 1997; 59. [PMID: 9074778]
  91. Gillespie PG., Walker RG. Molecular basis of mechanosensory transduction. Nature 2001; 413(6852). [PMID: 11557988]
  92.  Mechanotransduction involving multimodular proteins: converting force into biochemical signals. Annu Rev Biophys Biomol Struct 2006; 35. [PMID: 16689645]
  93. Svitkina TM., Verkhovsky AB., McQuade KM., Borisy GG. Analysis of the actin-myosin II system in fish epidermal keratocytes: mechanism of cell body translocation. J. Cell Biol. 1997; 139(2). [PMID: 9334344]
  94. Gupton SL., Eisenmann K., Alberts AS., Waterman-Storer CM. mDia2 regulates actin and focal adhesion dynamics and organization in the lamella for efficient epithelial cell migration. J. Cell. Sci. 2007; 120(Pt 19). [PMID: 17855386]
  95. Vicente-Manzanares M., Zareno J., Whitmore L., Choi CK., Horwitz AF. Regulation of protrusion, adhesion dynamics, and polarity by myosins IIA and IIB in migrating cells. J. Cell Biol. 2007; 176(5). [PMID: 17312025]
  96. Even-Ram S., Doyle AD., Conti MA., Matsumoto K., Adelstein RS., Yamada KM. Myosin IIA regulates cell motility and actomyosin-microtubule crosstalk. Nat. Cell Biol. 2007; 9(3). [PMID: 17310241]
  97. Riveline D., Zamir E., Balaban NQ., Schwarz US., Ishizaki T., Narumiya S., Kam Z., Geiger B., Bershadsky AD. Focal contacts as mechanosensors: externally applied local mechanical force induces growth of focal contacts by an mDia1-dependent and ROCK-independent mechanism. J. Cell Biol. 2001; 153(6). [PMID: 11402062]
  98. Giannone G., Dubin-Thaler BJ., Rossier O., Cai Y., Chaga O., Jiang G., Beaver W., Döbereiner HG., Freund Y., Borisy G., Sheetz MP. Lamellipodial actin mechanically links myosin activity with adhesion-site formation. Cell 2007; 128(3). [PMID: 17289574]
  99. McBeath R., Pirone DM., Nelson CM., Bhadriraju K., Chen CS. Cell shape, cytoskeletal tension, and RhoA regulate stem cell lineage commitment. Dev. Cell 2004; 6(4). [PMID: 15068789]
Updated on: Mon, 20 Oct 2014 09:18:17 GMT
History