The cytoskeleton and its components — actin, microtubules and intermediate filaments — have been studied for decades, and multiple roles of the individual cytoskeletal substructures are now well established. However, in recent years it has become apparent that the three cytoskeletal elements also engage in extensive crosstalk that is important for core biological processes. Actin–microtubule crosstalk is particularly important for the regulation of cell shape and polarity during cell migration and division and the establishment of neuronal and epithelial cell shape and function. This crosstalk engages different cytoskeletal regulators and encompasses various physical interactions, such as crosslinking, anchoring and mechanical support. Thus, the cytoskeleton should be considered not as a collection of individual parts but rather as a unified system in which subcomponents co-regulate each other to exert their functions in a precise and highly adaptable manner.
Access optionsAccess options
Subscribe to Journal
Get full journal access for 1 year
only $22.08 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Desai, A. & Mitchison, T. J. Microtubule polymerization dynamics. Annu. Rev. Cell Dev. Biol. 13, 83–117 (1997).
Blanchoin, L., Boujemaa-Paterski, R., Sykes, C. & Plastino, J. Actin dynamics, architecture, and mechanics in cell motility. Physiol. Rev. 94, 235–263 (2014).
Suozzi, K. C., Wu, X. & Fuchs, E. Spectraplakins: master orchestrators of cytoskeletal dynamics. J. Cell Biol. 197, 465–475 (2012).
Applewhite, D. A., Grode, K. D., Duncan, M. C. & Rogers, S. L. The actin-microtubule cross-linking activity of Drosophila Short stop is regulated by intramolecular inhibition. Mol. Biol. Cell 24, 2885–2893 (2013).
Preciado Lopez, M. et al. Actin-microtubule coordination at growing microtubule ends. Nat. Commun. 5, 4778 (2014). This paper reports the first in vitro reconstitution of actin–microtubule crosstalk mediated by crosslinking proteins localized to growing microtubule plus ends.
Janson, M. E., de Dood, M. E. & Dogterom, M. Dynamic instability of microtubules is regulated by force. J. Cell Biol. 161, 1029–1034 (2003).
Clark, A. G., Dierkes, K. & Paluch, E. K. Monitoring actin cortex thickness in live cells. Biophys. J. 105, 570–580 (2013).
Bradke, F. & Dotti, C. G. The role of local actin instability in axon formation. Science 283, 1931–1934 (1999).
Lancaster, O. M. et al. Mitotic rounding alters cell geometry to ensure efficient bipolar spindle formation. Dev. Cell 25, 270–283 (2013).
Luxenburg, C., Pasolli, H. A., Williams, S. E. & Fuchs, E. Developmental roles for Srf, cortical cytoskeleton and cell shape in epidermal spindle orientation. Nat. Cell Biol. 13, 203–214 (2011).
Chanet, S., Sharan, R., Khan, Z. & Martin, A. C. Myosin 2-induced mitotic rounding enables columnar epithelial cells to interpret cortical spindle positioning cues. Curr. Biol. 27, 3350–3358 (2017). This paper uses live-cell imaging of the first mitotic divisions of the early Drosophila melanogaster embryo to show that myosin-II-driven cell rounding is necessary for epithelial cells to orient their spindles in the plane of the epithelium instead of along the long apico-basal axis.
Basu, R. & Chang, F. Shaping the actin cytoskeleton using microtubule tips. Curr. Opin. Cell Biol. 19, 88–94 (2007).
Wen, Y. et al. EB1 and APC bind to mDia to stabilize microtubules downstream of Rho and promote cell migration. Nat. Cell Biol. 6, 820–830 (2004).
Okada, K. et al. Adenomatous polyposis coli protein nucleates actin assembly and synergizes with the formin mDia1. J. Cell Biol. 189, 1087–1096 (2010).
Henty-Ridilla, J. L., Rankova, A., Eskin, J. A., Kenny, K. & Goode, B. L. Accelerated actin filament polymerization from microtubule plus ends. Science 352, 1004–1009 (2016). This paper demonstrates the first in vitro reconstitution of actin filament polymerization from microtubule plus ends mediated by a complex of the microtubule +TIP protein CLIP170 and the actin elongation factor mDia1.
Lewkowicz, E. et al. The microtubule-binding protein CLIP-170 coordinates mDia1 and actin reorganization during CR3-mediated phagocytosis. J. Cell Biol. 183, 1287–1298 (2008).
Gaillard, J. et al. Differential interactions of the formins INF2, mDia1, and mDia2 with microtubules. Mol. Biol. Cell 22, 4575–4587 (2011).
Bartolini, F. et al. An mDia1-INF2 formin activation cascade facilitated by IQGAP1 regulates stable microtubules in migrating cells. Mol. Biol. Cell 27, 1797–1808 (2016).
Roth-Johnson, E. A., Vizcarra, C. L., Bois, J. S. & Quinlan, M. E. Interaction between microtubules and the Drosophila formin Cappuccino and its effect on actin assembly. J. Biol. Chem. 289, 4395–4404 (2014).
Szikora, S. et al. The formin DAAM is required for coordination of the actin and microtubule cytoskeleton in axonal growth cones. J. Cell Sci. 130, 2506–2519 (2017).
Wojnacki, J., Quassollo, G., Marzolo, M. P. & Caceres, A. Rho GTPases at the crossroad of signaling networks in mammals: impact of Rho-GTPases on microtubule organization and dynamics. Small GTPases 5, e28430 (2014).
Farina, F. et al. The centrosome is an actin-organizing centre. Nat. Cell Biol. 18, 65–75 (2016).
Dogterom, M. & Yurke, B. Measurement of the force-velocity relation for growing microtubules. Science 278, 856–860 (1997).
Ingber, D. E. Tensegrity, I. Cell structure and hierarchical systems biology. J. Cell Sci. 116, 1157–1173 (2003).
Brangwynne, C. P. et al. Microtubules can bear enhanced compressive loads in living cells because of lateral reinforcement. J. Cell Biol. 173, 733–741 (2006). This paper uses an elegant combination of quantitative live-cell imaging and theoretical modelling to explain how microtubules are reinforced against compressive loads owing to mechanical coupling to the surrounding (actin) cytoskeleton.
Bouchet, B. P. & Akhmanova, A. Microtubules in 3D cell motility. J. Cell Sci. 130, 39–50 (2017).
Lu, W., Fox, P., Lakonishok, M., Davidson, M. W. & Gelfand, V. I. Initial neurite outgrowth in Drosophila neurons is driven by kinesin-powered microtubule sliding. Curr. Biol. 23, 1018–1023 (2013).
Winding, M., Kelliher, M. T., Lu, W., Wildonger, J. & Gelfand, V. I. Role of kinesin-1-based microtubule sliding in Drosophila nervous system development. Proc. Natl Acad. Sci. USA 113, E4985–E4994 (2016).
Mogessie, B., Roth, D., Rahil, Z. & Straube, A. A novel isoform of MAP4 organises the paraxial microtubule array required for muscle cell differentiation. eLife 4, e05697 (2015).
Jolly, A. L. et al. Kinesin-1 heavy chain mediates microtubule sliding to drive changes in cell shape. Proc. Natl Acad. Sci. USA 107, 12151–12156 (2010).
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, 315–333 (2010).
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 Biol. 196, 363–374 (2012).
Paul, N. R., Jacquemet, G. & Caswell, P. T. Endocytic trafficking of integrins in cell migration. Curr. Biol. 25, R1092–R1105 (2015).
Schuler, M. H. et al. Miro1-mediated mitochondrial positioning shapes intracellular energy gradients required for cell migration. Mol. Biol. Cell 28, 2159–2169 (2017). This paper shows that microtubules serve to position mitochondria near the cell periphery, where they provide a localized source of energy to power actin-driven cell movements.
Etienne-Manneville, S. Microtubules in cell migration. Annu. Rev. Cell Dev. Biol. 29, 471–499 (2013).
Friedl, P. & Wolf, K. Plasticity of cell migration: a multiscale tuning model. J. Cell Biol. 188, 11–19 (2010).
Tomasek, J. J. & Hay, E. D. Analysis of the role of microfilaments and microtubules in acquisition of bipolarity and elongation of fibroblasts in hydrated collagen gels. J. Cell Biol. 99, 536–549 (1984).
Jayatilaka, H. et al. EB1 and cytoplasmic dynein mediate protrusion dynamics for efficient 3-dimensional cell migration. FASEB J. 32, 1207–1221 (2017).
Bouchet, B. P. et al. Mesenchymal cell invasion requires cooperative regulation of persistent microtubule growth by SLAIN2 and CLASP1. Dev. Cell 39, 708–723 (2016). This paper demonstrates that persistent microtubule growth maintained by the +TIP proteins SLAIN2 and CLASP1 is required for cell migration in 3D matrices but not on 2D surfaces.
Charras, G. & Sahai, E. Physical influences of the extracellular environment on cell migration. Nat. Rev. Mol. Cell Biol. 15, 813–824 (2014).
Huda, S. et al. Microtubule guidance tested through controlled cell geometry. J. Cell Sci. 125, 5790–5799 (2012).
Kaverina, I., Rottner, K. & Small, J. V. Targeting, capture, and stabilization of microtubules at early focal adhesions. J. Cell Biol. 142, 181–190 (1998).
Kaverina, I., Krylyshkina, O. & Small, J. V. Microtubule targeting of substrate contacts promotes their relaxation and dissociation. J. Cell Biol. 146, 1033–1044 (1999).
Krylyshkina, O. et al. Nanometer targeting of microtubules to focal adhesions. J. Cell Biol. 161, 853–859 (2003).
Stehbens, S. J. et al. CLASPs link focal-adhesion-associated microtubule capture to localized exocytosis and adhesion site turnover. Nat. Cell Biol. 16, 561–573 (2014).
Yu, X. et al. N-WASP coordinates the delivery and F-actin-mediated capture of MT1-MMP at invasive pseudopods. J. Cell Biol. 199, 527–544 (2012).
Rooney, C. et al. The Rac activator STEF (Tiam2) regulates cell migration by microtubule-mediated focal adhesion disassembly. EMBO Rep. 11, 292–298 (2010).
Nalbant, P., Chang, Y. C., Birkenfeld, J., Chang, Z. F. & Bokoch, G. M. Guanine nucleotide exchange factor-H1 regulates cell migration via localized activation of RhoA at the leading edge. Mol. Biol. Cell 20, 4070–4082 (2009).
Ezratty, E. J., Bertaux, C., Marcantonio, E. E. & Gundersen, G. G. Clathrin mediates integrin endocytosis for focal adhesion disassembly in migrating cells. J. Cell Biol. 187, 733–747 (2009).
Yue, J. et al. Microtubules regulate focal adhesion dynamics through MAP4K4. Dev. Cell 31, 572–585 (2014).
Ning, W. et al. The CAMSAP3-ACF7 complex couples noncentrosomal microtubules with actin filaments to coordinate their dynamics. Dev. Cell 39, 61–74 (2016).
Kodama, A., Karakesisoglou, I., Wong, E., Vaezi, A. & Fuchs, E. ACF7: an essential integrator of microtubule dynamics. Cell 115, 343–354 (2003).
Wu, X., Kodama, A. & Fuchs, E. ACF7 regulates cytoskeletal-focal adhesion dynamics and migration and has ATPase activity. Cell 135, 137–148 (2008). This pioneering study reports an essential function of actin–microtubule crosslinking by the spectraplakin ACF7 for directional cell migration in the skin epidermis.
Yue, J. et al. In vivo epidermal migration requires focal adhesion targeting of ACF7. Nat. Commun. 7, 11692 (2016).
Stroud, M. J. et al. GAS2-like proteins mediate communication between microtubules and actin through interactions with end-binding proteins. J. Cell Sci. 127, 2672–2682 (2014).
Jiang, K. et al. A Proteome-wide screen for mammalian SxIP motif-containing microtubule plus-end tracking proteins. Curr. Biol. 22, 1800–1807 (2012). This paper reports a proteome-wide search for mammalian +TIP proteins containing the core SxIP motif that recognizes EB proteins, revealing several new +TIPs that link microtubule plus ends to the actin cortex.
Girdler, G. C., Applewhite, D. A., Perry, W. M., Rogers, S. L. & Roper, K. The Gas2 family protein Pigs is a microtubule +TIP that affects cytoskeleton organisation. J. Cell Sci. 129, 121–134 (2016).
Bouchet, B. P. et al. Talin-KANK1 interaction controls the recruitment of cortical microtubule stabilizing complexes to focal adhesions. eLife 5, e18124 (2016). This paper identifies the molecular mechanisms that mediate the selective capture and stabilization of microtubules at the cell cortex in the vicinity of integrin-based adhesions to the ECM.
Byron, A. et al. A proteomic approach reveals integrin activation state-dependent control of microtubule cortical targeting. Nat. Commun. 6, 6135 (2015). This study uses an elegant combination of proteomics and live-cell imaging of microtubule–integrin interactions on micropatterned surfaces to demonstrate that active integrin complexes are specifically enriched for proteins associated with microtubule capture and stabilization.
Mimori-Kiyosue, Y. et al. CLASP1 and CLASP2 bind to EB1 and regulate microtubule plus-end dynamics at the cell cortex. J. Cell Biol. 168, 141–153 (2005).
Kumar, P. et al. GSK3beta phosphorylation modulates CLASP-microtubule association and lamella microtubule attachment. J. Cell Biol. 184, 895–908 (2009).
Zaoui, K., Benseddik, K., Daou, P., Salaun, D. & Badache, A. ErbB2 receptor controls microtubule capture by recruiting ACF7 to the plasma membrane of migrating cells. Proc. Natl Acad. Sci. USA 107, 18517–18522 (2010).
Margaron, Y., Fradet, N. & Cote, J. F. ELMO recruits actin cross-linking family 7 (ACF7) at the cell membrane for microtubule capture and stabilization of cellular protrusions. J. Biol. Chem. 288, 1184–1199 (2013).
Daou, P. et al. Essential and nonredundant roles for Diaphanous formins in cortical microtubule capture and directed cell migration. Mol. Biol. Cell 25, 658–668 (2014).
Lansbergen, G. et al. CLASPs attach microtubule plus ends to the cell cortex through a complex with LL5beta. Dev. Cell 11, 21–32 (2006).
Palazzo, A. F., Eng, C. H., Schlaepfer, D. D., Marcantonio, E. E. & Gundersen, G. G. Localized stabilization of microtubules by integrin- and FAK-facilitated Rho signaling. Science 303, 836–839 (2004).
Deakin, N. O. & Turner, C. E. Paxillin inhibits HDAC6 to regulate microtubule acetylation, Golgi structure, and polarized migration. J. Cell Biol. 206, 395–413 (2014).
Salmon, W. C., Adams, M. C. & Waterman-Storer, C. M. Dual-wavelength fluorescent speckle microscopy reveals coupling of microtubule and actin movements in migrating cells. J. Cell Biol. 158, 31–37 (2002).
Even-Ram, S. et al. Myosin IIA regulates cell motility and actomyosin-microtubule crosstalk. Nat. Cell Biol. 9, 299–309 (2007).
Waterman-Storer, C. M. & Salmon, E. D. 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, 417–434 (1997).
Gupton, S. L., Salmon, W. C. & Waterman-Storer, C. M. Converging populations of f-actin promote breakage of associated microtubules to spatially regulate microtubule turnover in migrating cells. Curr. Biol. 12, 1891–1899 (2002).
Mimori-Kiyosue, Y., Shiina, N. & Tsukita, S. Adenomatous polyposis coli (APC) protein moves along microtubules and concentrates at their growing ends in epithelial cells. J. Cell Biol. 148, 505–518 (2000).
Breitsprecher, D. et al. Rocket launcher mechanism of collaborative actin assembly defined by single-molecule imaging. Science 336, 1164–1168 (2012).
Campellone, K. G., Webb, N. J., Znameroski, E. A. & Welch, M. D. WHAMM is an Arp2/3 complex activator that binds microtubules and functions in ER to Golgi transport. Cell 134, 148–161 (2008).
Nejedla, M. et al. Profilin connects actin assembly with microtubule dynamics. Mol. Biol. Cell 27, 2381–2393 (2016).
Henty-Ridilla, J. L., Juanes, M. A. & Goode, B. L. Profilin directly promotes microtubule growth through residues mutated in amyotrophic lateral sclerosis. Curr. Biol. 27, 3535–3543 (2017).
Waterman-Storer, C. M., Worthylake, R. A., Liu, B. P., Burridge, K. & Salmon, E. D. Microtubule growth activates Rac1 to promote lamellipodial protrusion in fibroblasts. Nat. Cell Biol. 1, 45–50 (1999).
Lyle, K. S., Corleto, J. A. & Wittmann, T. Microtubule dynamics regulation contributes to endothelial morphogenesis. Bioarchitecture 2, 220–227 (2012).
Gierke, S. & Wittmann, T. EB1-recruited microtubule +TIP complexes coordinate protrusion dynamics during 3D epithelial remodeling. Curr. Biol. 22, 753–762 (2012).
Redd, M. J., Kelly, G., Dunn, G., Way, M. & Martin, P. Imaging macrophage chemotaxis in vivo: studies of microtubule function in zebrafish wound inflammation. Cell. Motil. Cytoskeleton 63, 415–422 (2006).
Yoo, S. K. et al. The role of microtubules in neutrophil polarity and migration in live zebrafish. J. Cell Sci. 125, 5702–5710 (2012).
Sugiyama, T., Pramanik, M. K. & Yumura, S. Microtubule-mediated inositol lipid signaling plays critical roles in regulation of blebbing. PLOS ONE 10, e0137032 (2015).
Conde, C. & Caceres, A. Microtubule assembly, organization and dynamics in axons and dendrites. Nat. Rev. Neurosci. 10, 319–332 (2009).
Xu, K., Zhong, G. & Zhuang, X. Actin, spectrin, and associated proteins form a periodic cytoskeletal structure in axons. Science 339, 452–456 (2013).
Shirao, T. & Gonzalez-Billault, C. Actin filaments and microtubules in dendritic spines. J. Neurochem. 126, 155–164 (2013).
Koser, D. E. et al. Mechanosensing is critical for axon growth in the developing brain. Nat. Neurosci. 19, 1592–1598 (2016).
Trivedi, N. et al. Drebrin-mediated microtubule-actomyosin coupling steers cerebellar granule neuron nucleokinesis and migration pathway selection. Nat. Commun. 8, 14484 (2017).
Coles, C. H. & Bradke, F. Coordinating neuronal actin-microtubule dynamics. Curr. Biol. 25, R677–R691 (2015).
Cammarata, G. M., Bearce, E. A. & Lowery, L. A. Cytoskeletal social networking in the growth cone: how +TIPs mediate microtubule-actin cross-linking to drive axon outgrowth and guidance. Cytoskeleton 73, 461–476 (2016).
Chia, J. X., Efimova, N. & Svitkina, T. M. Neurite outgrowth is driven by actin polymerization even in the presence of actin polymerization inhibitors. Mol. Biol. Cell 27, 3687–3790 (2016).
Geraldo, S., Khanzada, U. K., Parsons, M., Chilton, J. K. & Gordon-Weeks, P. R. Targeting of the F-actin-binding protein drebrin by the microtubule plus-tip protein EB3 is required for neuritogenesis. Nat. Cell Biol. 10, 1181–1189 (2008).
Elie, A. et al. Tau co-organizes dynamic microtubule and actin networks. Sci. Rep. 5, 9964 (2015).
Worth, D. C., Daly, C. N., Geraldo, S., Oozeer, F. & Gordon-Weeks, P. R. Drebrin contains a cryptic F-actin-bundling activity regulated by Cdk5 phosphorylation. J. Cell Biol. 202, 793–806 (2013).
van Haren, J. et al. Mammalian navigators are microtubule plus-end tracking proteins that can reorganize the cytoskeleton to induce neurite-like extensions. Cell. Motil. Cytoskeleton 66, 824–838 (2009).
Schmidt, K. L. et al. The cell migration molecule UNC-53/NAV2 is linked to the ARP2/3 complex by ABI-1. Development 136, 563–574 (2009).
Stringham, E. G. & Schmidt, K. L. Navigating the cell: UNC-53 and the navigators, a family of cytoskeletal regulators with multiple roles in cell migration, outgrowth and trafficking. Cell Adh. Migr. 3, 342–346 (2009).
Flynn, K. C. et al. ADF/cofilin-mediated actin retrograde flow directs neurite formation in the developing brain. Neuron 76, 1091–1107 (2012).
Flynn, K. C. The cytoskeleton and neurite initiation. Bioarchitecture 3, 86–109 (2013).
Witte, H., Neukirchen, D. & Bradke, F. Microtubule stabilization specifies initial neuronal polarization. J. Cell Biol. 180, 619–632 (2008).
Bradke, F. & Dotti, C. G. Neuronal polarity: vectorial cytoplasmic flow precedes axon formation. Neuron 19, 1175–1186 (1997).
Winans, A. M., Collins, S. R. & Meyer, T. Waves of actin and microtubule polymerization drive microtubule-based transport and neurite growth before single axon formation. eLife 5, e12387 (2016).
Zhou, F. Q., Waterman-Storer, C. M. & Cohan, C. S. Focal loss of actin bundles causes microtubule redistribution and growth cone turning. J. Cell Biol. 157, 839–849 (2002).
Burnette, D. T., Schaefer, A. W., Ji, L., Danuser, G. & Forscher, P. Filopodial actin bundles are not necessary for microtubule advance into the peripheral domain of Aplysia neuronal growth cones. Nat. Cell Biol. 9, 1360–1369 (2007).
Sanchez-Soriano, N. et al. Mouse ACF7 and drosophila short stop modulate filopodia formation and microtubule organisation during neuronal growth. J. Cell Sci. 122, 2534–2542 (2009).
Alves-Silva, J. et al. Spectraplakins promote microtubule-mediated axonal growth by functioning as structural microtubule-associated proteins and EB1-dependent +TIPs (tip interacting proteins). J. Neurosci. 32, 9143–9158 (2012). This paper establishes that actin–microtubule crosslinking at the lattice and the plus ends of microtubules by spectraplakins is essential for axonal outgrowth.
Ahmad, F. J. & Baas, P. W. Microtubules released from the neuronal centrosome are transported into the axon. J. Cell Sci. 108, 2761–2769 (1995).
del Castillo, U., Winding, M., Lu, W. & Gelfand, V. I. Interplay between kinesin-1 and cortical dynein during axonal outgrowth and microtubule organization in Drosophila neurons. eLife 4, e10140 (2015).
Grabham, P. W., Seale, G. E., Bennecib, M., Goldberg, D. J. & Vallee, R. B. Cytoplasmic dynein and LIS1 are required for microtubule advance during growth cone remodeling and fast axonal outgrowth. J. Neurosci. 27, 5823–5834 (2007).
Roossien, D. H., Lamoureux, P. & Miller, K. E. Cytoplasmic dynein pushes the cytoskeletal meshwork forward during axonal elongation. J. Cell Sci. 127, 3593–3602 (2014).
Ahmad, F. J. et al. Motor proteins regulate force interactions between microtubules and microfilaments in the axon. Nat. Cell Biol. 2, 276–280 (2000).
Bielas, S. L. et al. Spinophilin facilitates dephosphorylation of doublecortin by PP1 to mediate microtubule bundling at the axonal wrist. Cell 129, 579–591 (2007).
Burnette, D. T. et al. Myosin II activity facilitates microtubule bundling in the neuronal growth cone neck. Dev. Cell 15, 163–169 (2008).
Qu, Y., Hahn, I., Webb, S. E., Pearce, S. P. & Prokop, A. Periodic actin structures in neuronal axons are required to maintain microtubules. Mol. Biol. Cell 28, 296–308 (2017).
Krieg, M. et al. Genetic defects in beta-spectrin and tau sensitize C. elegans axons to movement-induced damage via torque-tension coupling. eLife 6, e20172 (2017). This paper reports an elegant combination of biophysical measurements and high-resolution microscopy in neurons with numerical simulations, showing how the spectrin and microtubule cytoskeletons work in concert to protect axons and dendrites from mechanical stress.
Fan, A., Tofangchi, A., Kandel, M., Popescu, G. & Saif, T. Coupled circumferential and axial tension driven by actin and myosin influences in vivo axon diameter. Sci. Rep. 7, 14188 (2017).
Jaworski, J. et al. Dynamic microtubules regulate dendritic spine morphology and synaptic plasticity. Neuron 61, 85–100 (2009).
Merriam, E. B. et al. Synaptic regulation of microtubule dynamics in dendritic spines by calcium, F-actin, and drebrin. J. Neurosci. 33, 16471–16482 (2013).
Rodriguez-Boulan, E. & Macara, I. G. Organization and execution of the epithelial polarity programme. Nat. Rev. Mol. Cell Biol. 15, 225–242 (2014).
Sugioka, K. & Sawa, H. Formation and functions of asymmetric microtubule organization in polarized cells. Curr. Opin. Cell Biol. 24, 517–525 (2012).
Reilein, A. & Nelson, W. J. APC is a component of an organizing template for cortical microtubule networks. Nat. Cell Biol. 7, 463–473 (2005).
Bazellieres, E. et al. Apico-basal elongation requires a drebrin-E-EB3 complex in columnar human epithelial cells. J. Cell Sci. 125, 919–931 (2012).
Gomez, J. M., Chumakova, L., Bulgakova, N. A. & Brown, N. H. Microtubule organization is determined by the shape of epithelial cells. Nat. Commun. 7, 13172 (2016).
Nashchekin, D., Fernandes, A. R. & St Johnston, D. Patronin/shot cortical foci assemble the noncentrosomal microtubule array that specifies the Drosophila anterior-posterior axis. Dev. Cell 38, 61–72 (2016).
Noordstra, I. et al. Control of apico-basal epithelial polarity by the microtubule minus-end-binding protein CAMSAP3 and spectraplakin ACF7. J. Cell Sci. 129, 4278–4288 (2016).
Toya, M. et al. CAMSAP3 orients the apical-to-basal polarity of microtubule arrays in epithelial cells. Proc. Natl Acad. Sci. USA 113, 332–337 (2016).
Hendershott, M. C. & Vale, R. D. Regulation of microtubule minus-end dynamics by CAMSAPs and Patronin. Proc. Natl Acad. Sci. USA 111, 5860–5865 (2014).
Jiang, K. et al. Microtubule minus-end stabilization by polymerization-driven CAMSAP deposition. Dev. Cell 28, 295–309 (2014).
Khanal, I., Elbediwy, A., Diaz de la Loza Mdel, C., Fletcher, G. C. & Thompson, B. J. Shot and Patronin polarise microtubules to direct membrane traffic and biogenesis of microvilli in epithelia. J. Cell Sci. 129, 2651–2659 (2016). This paper reveals the intricate bidirectional actin–microtubule crosstalk necessary to establish polarization of microtubules and actin microvilli along the apico-basal axis of epithelial cells.
Hotta, A. et al. Laminin-based cell adhesion anchors microtubule plus ends to the epithelial cell basal cortex through LL5alpha/beta. J. Cell Biol. 189, 901–917 (2010).
Chausovsky, A., Bershadsky, A. D. & Borisy, G. G. Cadherin-mediated regulation of microtubule dynamics. Nat. Cell Biol. 2, 797–804 (2000).
Waterman-Storer, C. M., Salmon, W. C. & Salmon, E. D. Feedback interactions between cell-cell adherens junctions and cytoskeletal dynamics in newt lung epithelial cells. Mol. Biol. Cell 11, 2471–2483 (2000).
Meng, W., Mushika, Y., Ichii, T. & Takeichi, M. Anchorage of microtubule minus ends to adherens junctions regulates epithelial cell-cell contacts. Cell 135, 948–959 (2008).
Ligon, L. A. & Holzbaur, E. L. Microtubules tethered at epithelial cell junctions by dynein facilitate efficient junction assembly. Traffic 8, 808–819 (2007).
Bellett, G. et al. Microtubule plus-end and minus-end capture at adherens junctions is involved in the assembly of apico-basal arrays in polarised epithelial cells. Cell. Motil. Cytoskeleton 66, 893–908 (2009).
Shaw, R. M. et al. Microtubule plus-end-tracking proteins target gap junctions directly from the cell interior to adherens junctions. Cell 128, 547–560 (2007).
Gavilan, M. P. et al. Alpha-catenin-dependent recruitment of the centrosomal protein CAP350 to adherens junctions allows epithelial cells to acquire a columnar shape. PLOS Biol. 13, e1002087 (2015).
Stehbens, S. J. et al. Dynamic microtubules regulate the local concentration of E-cadherin at cell-cell contacts. J. Cell Sci. 119, 1801–1811 (2006).
Shahbazi, M. N. et al. CLASP2 interacts with p120-catenin and governs microtubule dynamics at adherens junctions. J. Cell Biol. 203, 1043–1061 (2013).
Tsvetkov, A. S., Samsonov, A., Akhmanova, A., Galjart, N. & Popov, S. V. Microtubule-binding proteins CLASP1 and CLASP2 interact with actin filaments. Cell. Motil. Cytoskeleton 64, 519–530 (2007).
Karakesisoglou, I., Yang, Y. & Fuchs, E. An epidermal plakin that integrates actin and microtubule networks at cellular junctions. J. Cell Biol. 149, 195–208 (2000).
Mary, S. et al. Biogenesis of N-cadherin-dependent cell-cell contacts in living fibroblasts is a microtubule-dependent kinesin-driven mechanism. Mol. Biol. Cell 13, 285–301 (2002).
Sumigray, K. D., Foote, H. P. & Lechler, T. Noncentrosomal microtubules and type II myosins potentiate epidermal cell adhesion and barrier formation. J. Cell Biol. 199, 513–525 (2012).
Acharya, B. R. et al. KIF17 regulates RhoA-dependent actin remodeling at epithelial cell-cell adhesions. J. Cell Sci. 129, 957–970 (2016).
Komarova, Y. A. et al. VE-cadherin signaling induces EB3 phosphorylation to suppress microtubule growth and assemble adherens junctions. Mol. Cell 48, 914–925 (2012).
Plestant, C. et al. Adhesive interactions of N-cadherin limit the recruitment of microtubules to cell-cell contacts through organization of actomyosin. J. Cell Sci. 127, 1660–1671 (2014).
Holy, T. E., Dogterom, M., Yurke, B. & Leibler, S. Assembly and positioning of microtubule asters in microfabricated chambers. Proc. Natl Acad. Sci. USA 94, 6228–6231 (1997).
Laan, L. et al. Cortical dynein controls microtubule dynamics to generate pulling forces that position microtubule asters. Cell 148, 502–514 (2012).
Kimura, K. & Kimura, A. Intracellular organelles mediate cytoplasmic pulling force for centrosome centration in the Caenorhabditis elegans early embryo. Proc. Natl Acad. Sci. USA 108, 137–142 (2011).
Williams, S. E., Ratliff, L. A., Postiglione, M. P., Knoblich, J. A. & Fuchs, E. Par3-mInsc and Galphai3 cooperate to promote oriented epidermal cell divisions through LGN. Nat. Cell Biol. 16, 758–769 (2014).
Minc, N., Burgess, D. & Chang, F. Influence of cell geometry on division-plane positioning. Cell 144, 414–426 (2011).
Hertwig, O. (ed.) Das problem der Befruchtung and der Isotropie des Eies, eine Theorie der Vererbung 21–23 (Fischer, 1884).
Thery, M. et al. The extracellular matrix guides the orientation of the cell division axis. Nat. Cell Biol. 7, 947–953 (2005).
Sandquist, J. C., Kita, A. M. & Bement, W. M. And the dead shall rise: actin and myosin return to the spindle. Dev. Cell 21, 410–419 (2011).
di Pietro, F., Echard, A. & Morin, X. Regulation of mitotic spindle orientation: an integrated view. EMBO Rep. 17, 1106–1130 (2016).
Panousopoulou, E. & Green, J. B. Spindle orientation processes in epithelial growth and organisation. Semin. Cell Dev. Biol. 34, 124–132 (2014).
Holubcova, Z., Howard, G. & Schuh, M. Vesicles modulate an actin network for asymmetric spindle positioning. Nat. Cell Biol. 15, 937–947 (2013).
Mao, Y. et al. Planar polarization of the atypical myosin Dachs orients cell divisions in Drosophila. Genes Dev. 25, 131–136 (2011).
Machicoane, M. et al. SLK-dependent activation of ERMs controls LGN-NuMA localization and spindle orientation. J. Cell Biol. 205, 791–799 (2014).
Solinet, S. et al. The actin-binding ERM protein Moesin binds to and stabilizes microtubules at the cell cortex. J. Cell Biol. 202, 251–260 (2013).
Kunda, P. & Baum, B. The actin cytoskeleton in spindle assembly and positioning. Trends Cell Biol. 19, 174–179 (2009).
Maier, B., Kirsch, M., Anderhub, S., Zentgraf, H. & Kramer, A. The novel actin/focal adhesion-associated protein MISP is involved in mitotic spindle positioning in human cells. Cell Cycle 12, 1457–1471 (2013).
Zhu, M. et al. MISP is a novel Plk1 substrate required for proper spindle orientation and mitotic progression. J. Cell Biol. 200, 773–787 (2013).
Kwon, M., Bagonis, M., Danuser, G. & Pellman, D. Direct microtubule-binding by myosin-10 orients centrosomes toward retraction fibers and subcortical actin clouds. Dev. Cell 34, 323–337 (2015). This paper shows that myosin X localized at actin retraction fibres mediates centrosome positioning in dividing adherent cells, allowing the cell to retain a memory of its interphase adhesion pattern.
Fink, J. et al. External forces control mitotic spindle positioning. Nat. Cell Biol. 13, 771–778 (2011).
Matsumura, S. et al. Interphase adhesion geometry is transmitted to an internal regulator for spindle orientation via caveolin-1. Nat. Commun. 7, ncomms11858 (2016).
Bosveld, F. et al. Epithelial tricellular junctions act as interphase cell shape sensors to orient mitosis. Nature 530, 495–498 (2016).
Carminati, M. et al. Concomitant binding of Afadin to LGN and F-actin directs planar spindle orientation. Nat. Struct. Mol. Biol. 23, 155–163 (2016). This paper reports for the first time a mechanical anchor between dynein and the actin cortex in dividing epithelial cells that is based on the junctional protein afadin.
Gloerich, M., Bianchini, J. M., Siemers, K. A., Cohen, D. J. & Nelson, W. J. Cell division orientation is coupled to cell-cell adhesion by the E-cadherin/LGN complex. Nat. Commun. 8, 13996 (2017).
Redemann, S. et al. Membrane invaginations reveal cortical sites that pull on mitotic spindles in one-cell C. elegans embryos. PLOS ONE 5, e12301 (2010).
Woolner, S. & Papalopulu, N. Spindle position in symmetric cell divisions during epiboly is controlled by opposing and dynamic apicobasal forces. Dev. Cell 22, 775–787 (2012).
Bringmann, H. & Hyman, A. A. A cytokinesis furrow is positioned by two consecutive signals. Nature 436, 731–734 (2005).
Yuce, O., Piekny, A. & Glotzer, M. An ECT2-centralspindlin complex regulates the localization and function of RhoA. J. Cell Biol. 170, 571–582 (2005).
Canman, J. C. et al. Inhibition of Rac by the GAP activity of centralspindlin is essential for cytokinesis. Science 322, 1543–1546 (2008).
Foe, V. E. & von Dassow, G. Stable and dynamic microtubules coordinately shape the myosin activation zone during cytokinetic furrow formation. J. Cell Biol. 183, 457–470 (2008).
Fededa, J. P. & Gerlich, D. W. Molecular control of animal cell cytokinesis. Nat. Cell Biol. 14, 440–447 (2012).
Nguyen, P. A. et al. Spatial organization of cytokinesis signaling reconstituted in a cell-free system. Science 346, 244–247 (2014). This paper reports the first cell-free reconstitution of signalling between the mitotic spindle and the actomyosin contractile ring that occurs at the midplane of dividing cells using cytoplasmic frog egg extracts and model biomembranes.
Turlier, H., Audoly, B., Prost, J. & Joanny, J. F. Furrow constriction in animal cell cytokinesis. Biophys. J. 106, 114–123 (2014).
Gregory, S. L. et al. Cell division requires a direct link between microtubule-bound RacGAP and Anillin in the contractile ring. Curr. Biol. 18, 25–29 (2008).
van Oostende Triplet, C., Jaramillo Garcia, M., Haji Bik, H., Beaudet, D. & Piekny, A. Anillin interacts with microtubules and is part of the astral pathway that defines cortical domains. J. Cell Sci. 127, 3699–3710 (2014).
Tse, Y. C., Piekny, A. & Glotzer, M. Anillin promotes astral microtubule-directed cortical myosin polarization. Mol. Biol. Cell 22, 3165–3175 (2011).
Sampathkumar, A. et al. Live cell imaging reveals structural associations between the actin and microtubule cytoskeleton in Arabidopsis. Plant Cell 23, 2302–2313 (2011).
Hui, K. L. & Upadhyaya, A. Dynamic microtubules regulate cellular contractility during T cell activation. Proc. Natl Acad. Sci. USA 114, E4175–E4183 (2017).
Vleugel, M., Roth, S., Groenendijk, C. F. & Dogterom, M. Reconstitution of basic mitotic spindles in spherical emulsion droplets. J. Vis. Exp. 114, e54278 (2016).
Alvarado, J., Mulder, B. M. & Koenderink, G. H. Alignment of nematic and bundled semiflexible polymers in cell-sized confinement. Soft Matter 10, 2354–2364 (2014).
Tsai, F. C. & Koenderink, G. H. Shape control of lipid bilayer membranes by confined actin bundles. Soft Matter 11, 8834–8847 (2015).
Adikes, R. C., Hallett, R. A., Saway, B. F., Kuhlman, B. & Slep, K. C. Control of microtubule dynamics using an optogenetic microtubule plus end-F-actin cross-linker. J. Cell Biol. 217, 779–793 (2017).
Chen, B. C. et al. Lattice light-sheet microscopy: imaging molecules to embryos at high spatiotemporal resolution. Science 346, 1257998 (2014).
Welf, E. S. et al. Quantitative multiscale cell imaging in controlled 3D microenvironments. Dev. Cell 36, 462–475 (2016).
The authors thank A. Akhmanova for frequent discussions and insight. The authors thank C. Alkemade for preparing original schematics for all the figures. G.H.K. gratefully acknowledges support by the European Research Council (Starting Grant 335672-MINICELL). M.D. gratefully acknowledges support by the European Research Council (Synergy Grant 609822-MODEL CELL).
Nature Reviews Molecular Cell Biology thanks K. Slep and the other anonymous reviewer(s) for their contribution to the peer review of this work.
- Dynamic instability
A process of dynamic alternation between growing and shrinking states that is characteristic of microtubules and driven by the GTPase activity of tubulin.
- Cell cortex
A thin (~100 nm) filamentous meshwork of actin filaments and actin-binding proteins including myosin motors, which are tightly associated with the plasma membrane via proteins of the ezrin–radixin–moesin family. The cortex protects the mechanical integrity of the cell membrane and has a central role in cell shape control.
- Microtubule plus-end trackers
( +TIPs). Structurally diverse proteins that bind to the plus ends of growing microtubules. At least 20 different families of +TIPs exist. End-binding (EB) proteins are +TIPs that autonomously recognize growing microtubule ends. Other +TIPs bind to EB proteins through SxIP, Cap-Gly or LxxPTPh recognition motifs.+TIPs control microtubule dynamics and connect microtubules to various cellular structures, including the actin cortex, stress fibres and filopodial actin bundles.
The switch to rapid depolymerization triggered by the loss of the GTP cap at the growing end of the microtubule.
A sheet-like membrane protrusion that spans 2–4 µm from the leading edge of migrating and spreading cells and of neuronal growth cones. It contains a dense, branched network of actin filaments that polymerize at their plus ends near the leading edge and depolymerize at the back. The part of the leading edge directly behind the lamellipodium contains a more stable network of unbranched actin filaments and is enriched in myosin II.
- Focal adhesions
Adhesive junctions between cells and the ECM, which are mediated by integrins, whereby integrins interact with the ECM on the extracellular side and with actin bundles via adaptor and signalling proteins through their intracellular tails. Focal adhesions can contain over 100 different proteins, collectively referred to as the integrin adhesome. Cells modify the size and composition of focal adhesions in response to changes in the molecular composition and dimensionality (2D or 3D) of the matrix and physical forces.
- Leading edge
The front of a migrating cell. It is characterized by actin polymerization and the formation of nascent adhesions.
- Trailing edge
The rear end of a migrating cell. It is characterized by stable actin bundles and the release and disassembly of adhesions.
- Stress fibres
Bundles of 10–30 actin filaments crosslinked by α-actinin and often containing myosin II. There are four distinct types of stress fibre. Ventral stress fibres connect focal adhesions close to the cell edge to adhesions behind or near the nucleus. They are contractile and drive tail retraction and cell shape changes in migrating cells. Dorsal stress fibres are non-contractile but transmit contractile forces to the substrate via connections to focal adhesions. Transverse arcs are curved bundles behind the lamellipodium that are not connected to focal adhesions. They have been implicated in actin retrograde flow. The perinuclear actin cap is an ensemble of stress fibres that is anchored to the nucleus and controls its shape.
A type of membrane protrusion that contributes to the crawling-like cell migration of amoebas and of mammalian cells in 3D extracellular matrices. In white blood cells, pseudopodia enable the capture and engulfment of antigens. Pseudopodia are extended by the polymerization of a dense network of branched actin filaments at the leading edge and are supported by microtubules.
A process associated with the formation of blebs, which are round protrusions of the cell membrane caused by contraction of the actomyosin cortex in conjunction with a local rupture in the actin cortex or a transient detachment of the cortex from the cell membrane. Bleb expansion is driven by intracellular pressure generated in the cytoplasm whereas bleb retraction is driven by reformation of an actin cortex followed by myosin-driven contraction. Blebbing occurs during apoptosis, can drive the 3D motility of confined cells and acts as a pressure valve in dividing cells.
- Microtubule acetylation
A post-translational modification associated with long-lived microtubules whereby the Lys40 residue of α-tubulin in the microtubule lumen is enzymatically modified by tubulin acetyltransferase. Acetylation confers resilience against repeated mechanical stresses, thus protecting long-lived microtubules from mechanical ageing.
A regulatory protein that promotes actin assembly by sequestering monomeric actin, converting ADP-actin monomers into ATP-actin monomers and collaborating with actin nucleators such as formin to promote actin filament elongation.
Thin (60–200 nm) membrane protrusions that extend from the leading edge of lamellipodia in migrating cells, neuronal growth cones and epithelial sheets. They contain parallel bundles of 10–30 actin filaments crosslinked by fascin and fimbrin. Filopodia form focal adhesions with the substrate and sense the extracellular environment at their tips using cell surface receptors. In neurons, filopodia serve as precursors for dendrites.
- Navigator family
Microtubule-associated proteins that are expressed predominantly in the nervous system.
A family of actin-binding proteins that disassemble actin filaments by depolymerization at the minus end and by severing.
- Cytoplasmic flow
Refers to the movement of cytoplasm driven either by actomyosin contractility or by microtubule-based organelle movement. It is most common in plants and algae, but it also occurs during oogenesis in the fruitfly and during embryogenesis in Caenorhabditis elegans.
- Microtubule minus-end trackers
(–TIPs). Proteins that specifically bind to the minus end of non-centrosomal microtubules. The best-characterized proteins of the calmodulin-regulated spectrin-associated protein (CAMSAP)–Patronin–Nezha family protect minus ends from depolymerization and connect them to various cellular structures including the actin cortex at the apical surface of epithelial cells.
- Viscous drag
The frictional force that opposes the motion of an object in a viscous fluid. The viscous drag force is proportional to the velocity of the object, the fluid velocity and the object’s size, as expressed by Stokes’ law.
- Hertwig’s rule
A rule introduced by the German zoologist Oscar Hertwig in 1884 that is based on observations of the orientation of divisions of frog eggs upon controlled compression, stating that a cell divides along its long axis.
- Retraction fibres
Thin membrane tubes filled with actin filaments that maintain cell adhesion during mitotic rounding. They confer a memory of the cell–ECM adhesion geometry during interphase, allowing cells to orient their mitotic spindle.
- Planar cell division
Symmetrical cell division within the plane of an epithelial tissue. Planar alignment of the mitotic spindle is mediated by cortical cues, cell shape and mechanical tension. Coordinated planar cell divisions serve to elongate growing epithelial tissues while maintaining tissue cohesion.
A family of guanine nucleotide binding proteins present in the cell as hetero-oligomeric complexes. They form higher-order filamentous structures that can interact with actin, microtubules and membranes.