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10 - Microtubule Bending and Breaking in Cellular Mechanotransduction

Published online by Cambridge University Press:  05 July 2014

By
Andrew D. Bicek ,
Dominique Seetapun  and
David J. Odde
Edited by
Mohammad R. K. Mofrad  and
Roger D. Kamm
Andrew D. Bicek
Affiliation:
University of Minnesota
Dominique Seetapun
Affiliation:
University of Minnesota
David J. Odde
Affiliation:
University of Minnesota
Mohammad R. K. Mofrad
Affiliation:
University of California, Berkeley
Roger D. Kamm
Affiliation:
Massachusetts Institute of Technology
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Summary

Introduction

Cellular mechanotransduction is the mechanism by which living cells respond to mechanical signals from their environment. As early as 1892, Julius Wolff described the ability of bone to be deposited and resorbed in accordance with the mechanical stresses placed upon it, implying that the bone must have some internal mechanical stress or strain sensors (Huiskes and Verdonschot, 1997; Roesler, 1987; Wolff, 1892). More recently, investigating the precise biochemical mechanisms by which a direct mechanical stimulus is converted into a cellular response has become an area of interest, and the macro-scale effects of mechanotransduction, such as the alignment of load-bearing components, are now widely recognized. For example, the extracellular matrix protein, collagen, is organized into a hierarchy of fibrillar structures by tenocytes to form a tendon that functionally transmits mechanical tension (Kastelic et al., 1978). Additionally, vascular endothelial cells have been observed to align and alter their morphology in response to an applied fluid shear stress (Levesque and Nerem, 1985). In another example of cells sensing a mechanical stimulus, neuronal cells are capable of responding directly to a tensile force through neurite initiation and extension, a phenomenon termed “towed growth” (Bray, 1984; Fass and Odde, 2003; Fischer et al., 2005; Heidemann and Buxbaum, 1990; Pfister et al., 2004). Since individual cells are capable of responding directly to an applied force via secreting, organizing, and remodeling the extracellular matrix, or through morphological and gene expression changes, mechanotransduction is presumably controlled and integrated into a response at the cellular level.

Perhaps the best documentation of cellular mechanotransduction is the role of mechanically gated ion channels in hearing (Hudspeth, 1989). The stereocilia of the auditory hair cells vibrate and bend with incoming sound waves. As the stereocilia bend, a linker protein filament is tensed between two adjacent cilia and the tension generated opens a mechanically gated ion channel. Opening of the ion channel causes an influx of positive charges that depolarize the hair cell and lead to an electrical signal that the brain interprets as sound. While this is a clear example of a mechanotransduction event, it is also clear that mechanically gated ion channels are not the sole mechanism for mechanotransduction in every cell. Other structures within the cell therefore need to be identified and investigated for their mechanosensory features, with the cytoskeleton being a leading candidate.

Type
Chapter
Information
Cellular Mechanotransduction
Diverse Perspectives from Molecules to Tissues
 , pp. 234 - 249
Publisher: Cambridge University Press
Print publication year: 2009

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References

Ahmad, F. J., Yu, W., McNally, F. J. and Baas, P. W. (1999). An essential role for katanin in severing microtubules in the neuron. J Cell Biol 145, 305–15.CrossRefGoogle ScholarPubMed
Baas, P. W., Deitch, J. S., Black, M. M. and Banker, G. A. (1988). Polarity orientation of microtubules in hippocampal neurons: Uniformity in the axon and nonuniformity in the dendrite. Proc Natl Acad Sci USA 85, 8335–9.CrossRefGoogle ScholarPubMed
Baas, P. W., Karabay, A. and Qiang, L. (2005). Microtubules cut and run. Trends Cell Biol 15, 518–24.CrossRefGoogle Scholar
Baas, P. W., Vidya Nadar, C. and Myers, K. A. (2006). Axonal transport of microtubules: The long and short of it. Traffic 7, 490–8.CrossRefGoogle Scholar
Bicek, A. D., Tuzel, E., Demtchouk, A., Uppalapati, M., Hancock, W. O., Kroll, D. M., and Odde, D. J. (2009). Anterograde microtubule transport drives microtubule bending in LLC-PK1 epithelial cells. Mol Biol Cell 20, 2943–53.CrossRefGoogle ScholarPubMed
Brangwynne, C. P., MacKintosh, F. C., Kumar, S., Geisse, N. A., Talbot, J., Mahadevan, L., Parker, K. K., Ingber, D. E. and Weitz, D. A. (2006). Microtubules can bear enhanced compressive loads in living cells because of lateral reinforcement. J Cell Biol 173, 733–41.CrossRefGoogle ScholarPubMed
Bray, D. (1984). Axonal growth in response to experimentally applied mechanical tension. Dev Biol 102, 379–89.CrossRefGoogle ScholarPubMed
Briggs, M. W. and Sacks, D. B. (2003). IQGAP proteins are integral components of cytoskeletal regulation. EMBO Rep 4, 571–4.CrossRefGoogle ScholarPubMed
Brunner, D. and Nurse, P. (2000). CLIP170-like tip1p spatially organizes microtubular dynamics in fission yeast. Cell 102, 695–704.CrossRefGoogle ScholarPubMed
Buxbaum, R. E. and Heidemann, S. R. (1988). A thermodynamic model for force integration and microtubule assembly during axonal elongation. J Theor Biol 134, 379–90.CrossRefGoogle ScholarPubMed
Cassimeris, L., Gard, D., Tran, P. T. and Erickson, H. P. (2001). XMAP215 is a long thin molecule that does not increase microtubule stiffness. J Cell Sci 114, 3025–33.Google Scholar
Davis, L. J., Odde, D. J., Block, S. M. and Gross, S. P. (2002). The importance of lattice defects in katanin-mediated microtubule severing in vitro. Biophys J 82, 2916–27.CrossRefGoogle ScholarPubMed
Desai, A. and Mitchison, T. J. (1997). Microtubule polymerization dynamics. Annu Rev Cell Dev Biol 13, 83–117.CrossRefGoogle ScholarPubMed
Diamantopoulos, G. S., Perez, F., Goodson, H. V., Batelier, G., Melki, R., Kreis, T. E. and Rickard, J. E. (1999). Dynamic localization of CLIP-170 to microtubule plus ends is coupled to microtubule assembly. J Cell Biol 144, 99–112.CrossRefGoogle ScholarPubMed
Dillman, J. F., Dabney, L. P., Karki, S., Paschal, B. M., Holzbaur, E. L. and Pfister, K. K. (1996). Functional analysis of dynactin and cytoplasmic dynein in slow axonal transport. J Neurosci 16, 6742–52.CrossRefGoogle ScholarPubMed
Dogterom, M. and Yurke, B. (1997). Measurement of the force-velocity relation for growing microtubules. Science 278, 856–60.CrossRefGoogle ScholarPubMed
Dotti, C. G., Sullivan, C. A. and Banker, G. A. (1988). The establishment of polarity by hippocampal neurons in culture. J Neurosci 8, 1454–68.CrossRefGoogle ScholarPubMed
Drummond, D. R. and Cross, R. A. (2000). Dynamics of interphase microtubules in Schizosaccharomyces pombe. Curr Biol 10, 766–75.CrossRefGoogle ScholarPubMed
Errico, A., Ballabio, A. and Rugarli, E. I. (2002). Spastin, the protein mutated in autosomal dominant hereditary spastic paraplegia, is involved in microtubule dynamics. Hum Mol Genet 11, 153–63.CrossRefGoogle ScholarPubMed
Etienne-Manneville, S. and Hall, A. (2003). Cell polarity: Par6, aPKC and cytoskeletal crosstalk. Curr Opin Cell Biol 15, 67–72.CrossRefGoogle ScholarPubMed
Evans, K. J., Gomes, E. R., Reisenweber, S. M., Gundersen, G. G. and Lauring, B. P. (2005). Linking axonal degeneration to microtubule remodeling by Spastin-mediated microtubule severing. J Cell Biol 168, 599–606.CrossRefGoogle ScholarPubMed
Fass, J. N. and Odde, D. J. (2003). Tensile force-dependent neurite elicitation via anti-beta1 integrin antibody-coated magnetic beads. Biophys J 85, 623–36.CrossRefGoogle ScholarPubMed
Felgner, H., Frank, R., Biernat, J., Mandelkow, E. M., Mandelkow, E., Ludin, B., Matus, A. and Schliwa, M. (1997). Domains of neuronal microtubule-associated proteins and flexural rigidity of microtubules. J Cell Biol 138, 1067–75.CrossRefGoogle ScholarPubMed
Felgner, H., Frank, R. and Schliwa, M. (1996). Flexural rigidity of microtubules measured with the use of optical tweezers. J Cell Sci 109(Pt 2), 509–16.Google ScholarPubMed
Fischer, T. M., Steinmetz, P. N. and Odde, D. J. (2005). Robust micromechanical neurite elicitation in synapse-competent neurons via magnetic bead force application. Ann Biomed Eng 33, 1229–37.CrossRefGoogle ScholarPubMed
Forscher, P. and Smith, S. J. (1988). Actions of cytochalasins on the organization of actin filaments and microtubules in a neuronal growth cone. J Cell Biol 107, 1505–16.CrossRefGoogle Scholar
Fukata, M., Nakagawa, M. and Kaibuchi, K. (2003). Roles of Rho-family GTPases in cell polarisation and directional migration. Curr Opin Cell Biol 15, 590–7.CrossRefGoogle ScholarPubMed
Fukata, M., Watanabe, T., Noritake, J., Nakagawa, M., Yamaga, M., Kuroda, S., Matsuura, Y., Iwamatsu, A., Perez, F. and Kaibuchi, K. (2002). Rac1 and Cdc42 capture microtubules through IQGAP1 and CLIP-170. Cell 109, 873–85.CrossRefGoogle ScholarPubMed
Fygenson, D. K., Elbaum, M., Shraiman, B. and Libchaber, A. (1997). Microtubules and vesicles under controlled tension. Phys Rev E 55, 850–59.CrossRefGoogle Scholar
Gittes, F., Mickey, B., Nettleton, J. and Howard, J. (1993). Flexural rigidity of microtubules and actin filaments measured from thermal fluctuations in shape. J Cell Biol 120, 923–34.CrossRefGoogle Scholar
Goldberg, D. J. and Burmeister, D. W. (1986). Stages in axon formation: Observations of growth of Aplysia axons in culture using video-enhanced contrast-differential interference contrast microscopy. J Cell Biol 103, 1921–31.CrossRefGoogle ScholarPubMed
Gundersen, G. G. and Bulinski, J. C. (1988). Selective stabilization of microtubules oriented toward the direction of cell migration. Proc Natl Acad Sci USA 85, 5946–50.CrossRefGoogle ScholarPubMed
Gupton, S. L., Salmon, W. C. and Waterman-Storer, C. M. (2002). Converging populations of f-actin promote breakage of associated microtubules to spatially regulate microtubule turnover in migrating cells. Curr Biol 12, 1891–9.CrossRefGoogle ScholarPubMed
Heidemann, S. R. and Buxbaum, R. E. (1990). Tension as a regulator and integrator of axonal growth. Cell Motil Cytoskeleton 17, 6–10.CrossRefGoogle ScholarPubMed
Heidemann, S. R., Kaech, S., Buxbaum, R. E. and Matus, A. (1999). Direct observations of the mechanical behaviors of the cytoskeleton in living fibroblasts. J Cell Biol 145, 109–22.CrossRefGoogle ScholarPubMed
Howard, J. (2001). Mechanics of Motor Proteins and the Cytoskeleton. Sunderland, MA: Sinauer Associates.Google Scholar
Hudspeth, A. J. (1989). How the ear’s works work. Nature 341, 397–404.CrossRefGoogle ScholarPubMed
Huiskes, R. and Verdonschot, N. (1997). Biomechanics of Artificial Joints: The Hip. In Basic Orthopaedic Biomechanics, 2nd ed. (ed. Mow, V. C. and Hayes, W. C.). Philadelphia: Lippincott-Raven Publishers.Google Scholar
Ingber, D. E. (1993). Cellular tensegrity: Defining new rules of biological design that govern the cytoskeleton. J Cell Sci 104(Pt 3), 613–27.Google ScholarPubMed
Janson, M. E., de Dood, M. E. and Dogterom, M. (2003). Dynamic instability of microtubules is regulated by force. J Cell Biol 161, 1029–34.CrossRefGoogle ScholarPubMed
Janson, M. E. and Dogterom, M. (2004). A bending mode analysis for growing microtubules: Evidence for a velocity-dependent rigidity. Biophys J 87, 2723–36.CrossRefGoogle ScholarPubMed
Kastelic, J., Galeski, A. and Baer, E. (1978). The multicomposite structure of tendon. Connect Tissue Res 6, 11–23.CrossRefGoogle ScholarPubMed
Kaverina, I., Krylyshkina, O., Beningo, K., Anderson, K., Wang, Y. L. and Small, J. V. (2002). Tensile stress stimulates microtubule outgrowth in living cells. J Cell Sci 115, 2283–91.Google ScholarPubMed
Keating, T. J., Peloquin, J. G., Rodionov, V. I., Momcilovic, D. and Borisy, G. G. (1997). Microtubule release from the centrosome. Proc Natl Acad Sci USA 94, 5078–83.CrossRefGoogle ScholarPubMed
Khachigian, L. M., Resnick, N., Gimbrone, M. A. and Collins, T. (1995). Nuclear factor-kappa B interacts functionally with the platelet-derived growth factor B-chain shear-stress response element in vascular endothelial cells exposed to fluid shear stress. J Clin Invest 96, 1169–75.CrossRefGoogle ScholarPubMed
Kirschner, M. W. and Mitchison, T. (1986). Microtubule dynamics. Nature 324, 621.CrossRefGoogle ScholarPubMed
Kis, A., Kasas, S., Babic, B., Kulik, A. J., Benoit, W., Briggs, G. A. D., Schonenberger, C., Catsicas, S. and Forro, L. (2002). Nanomechanics of microtubules. Phys Rev Lett 89, 248101.CrossRefGoogle ScholarPubMed
Komarova, Y. A., Akhmanova, A. S., Kojima, S., Galjart, N. and Borisy, G. G. (2002). Cytoplasmic linker proteins promote microtubule rescue in vivo. J Cell Biol 159, 589–99.CrossRefGoogle ScholarPubMed
Kurachi, M., Hoshi, M. and Tashiro, H. (1995). Buckling of a single microtubule by optical trapping forces: Direct measurement of microtubule rigidity. Cell Motil Cytoskeleton 30, 221–8.CrossRefGoogle ScholarPubMed
Kurz, J. C. and Williams, R. C.. (1995). Microtubule-associated proteins and the flexibility of microtubules. Biochemistry 34, 13374–80.CrossRefGoogle ScholarPubMed
Levesque, M. J. and Nerem, R. M. (1985). The elongation and orientation of cultured endothelial cells in response to shear stress. J Biomech Eng 107, 341–7.CrossRefGoogle ScholarPubMed
Liao, G., Nagasaki, T. and Gundersen, G. G. (1995). Low concentrations of nocodazole interfere with fibroblast locomotion without significantly affecting microtubule level: Implications for the role of dynamic microtubules in cell locomotion. J Cell Sci 108, 3473–83.Google ScholarPubMed
Lohret, T. A., Zhao, L. and Quarmby, L. M. (1999). Cloning of Chlamydomonas p60 katanin and localization to the site of outer doublet severing during deflagellation. Cell Motil Cytoskeleton 43, 221–31.3.0.CO;2-E>CrossRefGoogle ScholarPubMed
McNally, F. J. (2001). Cytoskeleton: CLASPing the end to the edge. Curr Biol 11, R477–80.CrossRefGoogle Scholar
McNally, F. J. and Thomas, S. (1998). Katanin is responsible for the M-phase microtubule-severing activity in Xenopus eggs. Mol Biol Cell 9, 1847–61.CrossRefGoogle ScholarPubMed
McNally, F. J. and Vale, R. D. (1993). Identification of katanin, an ATPase that severs and disassembles stable microtubules. Cell 75, 419–29.CrossRefGoogle ScholarPubMed
McNally, K., Audhya, A., Oegema, K. and McNally, F. J. (2006). Katanin controls mitotic and meiotic spindle length. J Cell Biol 175, 881–91.CrossRefGoogle ScholarPubMed
McNally, K. P., Buster, D. and McNally, F. J. (2002). Katanin-mediated microtubule severing can be regulated by multiple mechanisms. Cell Motil Cytoskeleton 53, 337–49.CrossRefGoogle ScholarPubMed
Mickey, B. and Howard, J. (1995). Rigidity of microtubules is increased by stabilizing agents. J Cell Biol 130, 909–17.CrossRefGoogle ScholarPubMed
Mitchison, T. and Kirschner, M. (1984). Dynamic instability of microtubule growth. Nature 312, 237–42.CrossRefGoogle ScholarPubMed
Mitchison, T. J. (1993). Localization of an exchangeable GTP binding site at the plus end of microtubules. Science 261, 1044–7.CrossRefGoogle ScholarPubMed
Nagasaki, T., Chapin, C. J. and Gundersen, G. G. (1992). Distribution of detyrosinated microtubules in motile NRK fibroblasts is rapidly altered upon cell-cell contact: Implications for contact inhibition of locomotion. Cell Motil Cytoskeleton 23, 45–60.CrossRefGoogle ScholarPubMed
Nishimura, T., Kato, K., Yamaguchi, T., Fukata, Y., Ohno, S. and Kaibuchi, K. (2004). Role of the PAR-3-KIF3 complex in the establishment of neuronal polarity. Nat Cell Biol 6, 328–34.CrossRefGoogle ScholarPubMed
Odde, D. J., Ma, L., Briggs, A. H., DeMarco, A. and Kirschner, M. W. (1999). Microtubule bending and breaking in living fibroblast cells. J Cell Sci 112(Pt 19), 3283–8.Google ScholarPubMed
Pampaloni, F., Lattanzi, G., Jonas, A., Surrey, T., Frey, E. and Florin, E. L. (2006). Thermal fluctuations of grafted microtubules provide evidence of a length-dependent persistence length. Proc Natl Acad Sci USA 103, 10248–53.CrossRefGoogle ScholarPubMed
Perez, F., Diamantopoulos, G. S., Stalder, R. and Kreis, T. E. (1999). CLIP-170 highlights growing microtubule ends in vivo. Cell 96, 517–27.CrossRefGoogle ScholarPubMed
Pfister, B. J., Iwata, A., Meaney, D. F. and Smith, D. H. (2004). Extreme stretch growth of integrated axons. J Neurosci 24, 7978–83.CrossRefGoogle ScholarPubMed
Pfister, K. K. (1999). Cytoplasmic dynein and microtubule transport in the axon: The action connection. Mol Neurobiol 20, 81–91.CrossRefGoogle ScholarPubMed
Putnam, A. J., Cunningham, J. J., Dennis, R. G., Linderman, J. J. and Mooney, D. J. (1998). Microtubule assembly is regulated by externally applied strain in cultured smooth muscle cells. J Cell Sci 111(Pt 22), 3379–87.Google ScholarPubMed
Putnam, A. J., Cunningham, J. J., Pillemer, B. B. and Mooney, D. J. (2003). External mechanical strain regulates membrane targeting of Rho GTPases by controlling microtubule assembly. Am J Physiol Cell Physiol 284, C627–39.CrossRefGoogle ScholarPubMed
Putnam, A. J., Schultz, K. and Mooney, D. J. (2001). Control of microtubule assembly by extracellular matrix and externally applied strain. Am J Physiol Cell Physiol 280, C556–64.CrossRefGoogle ScholarPubMed
Ren, X. D., Kiosses, W. B. and Schwartz, M. A. (1999). Regulation of the small GTP-binding protein Rho by cell adhesion and the cytoskeleton. Embo J 18, 578–85.CrossRefGoogle Scholar
Roesler, H. (1987). The history of some fundamental concepts in bone biomechanics. J Biomech 20, 1025–34.CrossRefGoogle ScholarPubMed
Rosette, C. and Karin, M. (1995). Cytoskeletal control of gene expression: Depolymerization of microtubules activates NF-kappa B. J Cell Biol 128, 1111–9.CrossRefGoogle ScholarPubMed
Salmon, W. C., Adams, M. C. and Waterman-Storer, C. M. (2002). Dual-wavelength fluorescent speckle microscopy reveals coupling of microtubule and actin movements in migrating cells. J Cell Biol 158, 31–7.CrossRefGoogle ScholarPubMed
Schaefer, A. W., Kabir, N. and Forscher, P. (2002). Filopodia and actin arcs guide the assembly and transport of two populations of microtubules with unique dynamic parameters in neuronal growth cones. J Cell Biol 158, 139–52.CrossRefGoogle ScholarPubMed
Schneider, S. Q. and Bowerman, B. (2003). Cell polarity and the cytoskeleton in the Caenorhabditis elegans zygote. Annu Rev Genet 37, 221–49.CrossRefGoogle ScholarPubMed
Shi, S. H., Cheng, T., Jan, L. Y. and Jan, Y. N. (2004). APC and GSK-3beta are involved in mPar3 targeting to the nascent axon and establishment of neuronal polarity. Curr Biol 14, 2025–32.CrossRefGoogle ScholarPubMed
Shi, S. H., Jan, L. Y. and Jan, Y. N. (2003). Hippocampal neuronal polarity specified by spatially localized mPar3/mPar6 and PI 3-kinase activity. Cell 112, 63–75.CrossRefGoogle ScholarPubMed
Siegrist, S. E. and Doe, C. Q. (2007). Microtubule-induced cortical cell polarity. Genes Dev 21, 483–96.CrossRefGoogle ScholarPubMed
Stamenovic, D., Mijailovich, S. M., Tolic-Norrelykke, I. M., Chen, J. and Wang, N. (2001). Cell prestress. II. Contribution of microtubules. Am J Physiol Cell Physiol 282, C617–C624.CrossRefGoogle Scholar
Suter, D. M. and Forscher, P. (1998). An emerging link between cytoskeletal dynamics and cell adhesion molecules in growth cone guidance. Curr Opin Neurobiol 8, 106–16.CrossRefGoogle Scholar
Takasone, T., Juodkazis, S., Kawagishi, Y., Yamaguchi, A., Matsuo, S., Sakakibara, H., Nakayama, H. and Misawa., H. (2002). Flexural rigidity of a single microtubule. Jpn J Appl Phys 41, 3015–19.CrossRefGoogle Scholar
Tran, P. T., Marsh, L., Doye, V., Inoue, S. and Chang, F. (2001). A mechanism for nuclear positioning in fission yeast based on microtubule pushing. J Cell Biol 153, 397–411.CrossRefGoogle ScholarPubMed
Vale, R. D. (1991). Severing of stable microtubules by a mitotically activated protein in Xenopus egg extracts. Cell 64, 827–39.CrossRefGoogle ScholarPubMed
Vale, R. D., Reese, T. S. and Sheetz, M. P. (1985). Identification of a novel force-generating protein, kinesin, involved in microtubule-based motility. Cell 42, 39–50.CrossRefGoogle ScholarPubMed
Vallee, R. B., Wall, J. S., Paschal, B. M. and Shpetner, H. S. (1988). Microtubule-associated protein 1C from brain is a two-headed cytosolic dynein. Nature 332, 561–3.CrossRefGoogle Scholar
Venier, P., Maggs, A. C., Carlier, M. F. and Pantaloni, D. (1994). Analysis of microtubule rigidity using hydrodynamic flow and thermal fluctuations. J Biol Chem 269, 13353–60.Google ScholarPubMed
Vorobjev, I. A., Rodionov, V. I., Maly, I. V. and Borisy, G. G. (1999). Contribution of plus and minus end pathways to microtubule turnover. J Cell Sci 112(Pt 14), 2277–89.Google ScholarPubMed
Walker, R. A., O’Brien, E. T., Pryer, N. K., Soboeiro, M. F., Voter, W. A., Erickson, H. P. and Salmon, E. D. (1988). Dynamic instability of individual microtubules analyzed by video light microscopy: Rate constants and transition frequencies. J Cell Biol 107, 1437–48.CrossRefGoogle ScholarPubMed
Wang, L. and Brown, A. (2002). Rapid movement of microtubules in axons. Curr Biol 12, 1496–501.CrossRefGoogle ScholarPubMed
Wang, N., Butler, J. P. and Ingber, D. E. (1993). Mechanotransduction across the cell surface and through the cytoskeleton. Science 260, 1124–7.CrossRefGoogle ScholarPubMed
Wang, N., Naruse, K., Stamenovic, D., Fredberg, J. J., Mijailovich, S. M., Tolic-Norrelykke, I. M., Polte, T., Mannix, R. and Ingber, D. E. (2001). Mechanical behavior in living cells consistent with the tensegrity model. Proc Natl Acad Sci USA 98, 7765–70.CrossRefGoogle ScholarPubMed
Watanabe, T., Wang, S., Noritake, J., Sato, K., Fukata, M., Takefuji, M., Nakagawa, M., Izumi, N., Akiyama, T. and Kaibuchi, K. (2004). Interaction with IQGAP1 links APC to Rac1, Cdc42, and actin filaments during cell polarization and migration. Dev Cell 7, 871–83.CrossRefGoogle ScholarPubMed
Waterman-Storer, C. M. and Salmon, E. D. (1997). Actomyosin-based retrograde flow of microtubules in the lamella of migrating epithelial cells influences microtubule dynamic instability and turnover and is associated with microtubule breakage and treadmilling. J Cell Biol 139, 417–34.CrossRefGoogle ScholarPubMed
Waterman-Storer, C. M., Worthylake, R. A., Liu, B. P., Burridge, K. and Salmon, E. D. (1999). Microtubule growth activates Rac1 to promote lamellipodial protrusion in fibroblasts. Nat Cell Biol 1, 45–50.CrossRefGoogle ScholarPubMed
Wittmann, T. and Waterman-Storer, C. M. (2001). Cell motility: Can Rho GTPases and microtubules point the way?J Cell Sci 114, 3795–803.Google Scholar
Wolff, J. (1892). Das Gesetz der Transformation der knochen [The Law of Bone Remodeling]. Berlin: A. Hirschwald.Google Scholar
Yang, H. Y., McNally, K. and McNally, F. J. (2003). MEI-1/katanin is required for translocation of the meiosis I spindle to the oocyte cortex in C elegans. Dev Biol 260, 245–59.CrossRefGoogle ScholarPubMed
Yu, W., Ahmad, F. J. and Baas, P. W. (1994). Microtubule fragmentation and partitioning in the axon during collateral branch formation. J Neurosci 14, 5872–84.CrossRefGoogle ScholarPubMed
Zhou, F. Q., Waterman-Storer, C. M. and Cohan, C. S. (2002). Focal loss of actin bundles causes microtubule redistribution and growth cone turning. J Cell Biol 157, 839–49.CrossRefGoogle ScholarPubMed

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