Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Article
  • Published:

Competition for actin between two distinct F-actin networks defines a bistable switch for cell polarization

Abstract

Symmetry-breaking polarization enables functional plasticity of cells and tissues and is yet not well understood. Here we show that epithelial cells, hard-wired to maintain a static morphology and to preserve tissue organization, can spontaneously switch to a migratory polarized phenotype after relaxation of the actomyosin cytoskeleton. We find that myosin II engages actin in the formation of cortical actomyosin bundles and thus makes it unavailable for deployment in the process of dendritic growth normally driving cell motility. Under low-contractility regimes, epithelial cells polarize in a front–back manner owing to the emergence of actin retrograde flows powered by dendritic polymerization of actin. Coupled to cell movement, the flows transport myosin II from the front to the back of the cell, where the motor locally ‘locks’ actin in contractile bundles. This polarization mechanism could be employed by embryonic and cancer epithelial cells in microenvironments where high-contractility-driven cell motion is inefficient.

This is a preview of subscription content, access via your institution

Access options

Rent or buy this article

Prices vary by article type

from$1.95

to$39.95

Prices may be subject to local taxes which are calculated during checkout

Figure 1: Acute inhibition of myosin II activity results in spontaneous symmetry breaking and motility initiation in single epithelial cells.
Figure 2: Modulating cell–substrate adhesion strength per se is insufficient to trigger spontaneous symmetry breaking and motility initiation in epithelial cells with intact myosin II activity.
Figure 3: Myosin II activity suppresses actin polymerization at the cell edge by immobilizing actin inside peripheral actomyosin bundles.
Figure 4: The growth of submembranous actin at the edge of myosin II-inhibited cells is strongly reinforced through Arp2/3-mediated F-actin branching.
Figure 5: Mathematical model for migratory cell polarization based on the mass balance between actomyosin bundle formation and branched actin growth coupled to F-actin retrograde flow.
Figure 6: Low-contractility regimes enable spatial segregation among branched and bundled actin networks and facilitate actin–myosin flows.
Figure 7: Experimental validation of model-predicted core motifs that compose the actin/myosin polarity-generating system.
Figure 8: Testing generality of the actin/myosin polarity-generating mechanism in the context of epithelial biology.

Similar content being viewed by others

References

  1. Nieto, M. A. Epithelial plasticity: a common theme in embryonic and cancer cells. Science 342, 1234850 (2013).

    Article  Google Scholar 

  2. Muthuswamy, S. K. & Xue, B. Cell polarity as a regulator of cancer cell behavior plasticity. Annu. Rev. Cell Dev. Biol. 28, 599–625 (2012).

    Article  CAS  Google Scholar 

  3. Hall, A. Rho GTPases and the actin cytoskeleton. Science 279, 509–514 (1998).

    Article  CAS  Google Scholar 

  4. Bakal, C., Aach, J., Church, G. & Perrimon, N. Quantitative morphological signatures define local signaling networks regulating cell morphology. Science 316, 1753–1756 (2007).

    Article  CAS  Google Scholar 

  5. Sahai, E. & Marshall, C. J. Differing modes of tumour cell invasion have distinct requirements for Rho/ROCK signalling and extracellular proteolysis. Nat. Cell Biol. 8, 711–719 (2003).

    Article  Google Scholar 

  6. Sanz-Moreno, V. et al. Rac activation and inactivation control plasticity of tumor cell movement. Cell 135, 510–523 (2008).

    Article  CAS  Google Scholar 

  7. Croft, D. R. & Olson, M. F. Regulating the conversion between rounded and elongated modes of cancer cell movement. Cancer Cell 14, 349–351 (2008).

    Article  CAS  Google Scholar 

  8. Even-Ram, S. et al. Myosin IIA regulates cell motility and actomyosin-microtubule crosstalk. Nat. Cell Biol. 3, 299–309 (2007).

    Article  Google Scholar 

  9. Schramek, D. et al. Direct in vivo RNAi screen unveils myosin IIa as a tumor suppressor of squamous cell carcinomas. Science 343, 309–313 (2014).

    Article  CAS  Google Scholar 

  10. Vicente-Manzanares, M., Zareno, J., Whitmore, L., Choi, C. K. & Horwitz, A. F. Regulation of protrusion, adhesion dynamics, and polarity by myosins IIA and IIB in migrating cells. J. Cell Biol. 176, 573–580 (2007).

    Article  CAS  Google Scholar 

  11. Vicente-Manzanares, M., Ma, X., Adelstein, R. S. & Horwitz, A. R. Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat. Rev. Mol. Cell Biol. 10, 778–790 (2009).

    Article  CAS  Google Scholar 

  12. Gupton, S. L. & Waterman-Storer, C. M. Spatiotemporal feedback between actomyosin and focal-adhesion systems optimizes rapid cell migration. Cell 125, 1361–1374 (2006).

    Article  CAS  Google Scholar 

  13. Wu, X., Kodama, A. & Fuchs, E. ACF7 regulates cytoskeletal-focal adhesion dynamics and migration and has ATPase activity. Cell 135, 137–148 (2008).

    Article  CAS  Google Scholar 

  14. Barnhart, E., Lee, K. C., Allen, G. M., Theriot, J. A. & Mogilner, A. Balance between cell-substrate adhesion and myosin contraction determines the frequency of motility initiation in fish keratocytes. Proc. Natl Acad. Sci. USA 112, 5045–5050 (2015).

    Article  CAS  Google Scholar 

  15. Liu, Y. J. et al. Confinement and low adhesion induce fast amoeboid migration of slow mesenchymal cells. Cell 160, 659–672 (2015).

    Article  CAS  Google Scholar 

  16. Watanabe, T., Hosoya, H. & Yonemura, S. Regulation of myosin II dynamics by phosphorylation and dephosphorylation of its light chain in epithelial cells. Mol. Biol. Cell 18, 605–616 (2007).

    Article  CAS  Google Scholar 

  17. Cramer, L. P., Briggs, L. J. & Dawe, H. R. Use of fluorescently labelled deoxyribonuclease I to spatially measure G-actin levels in migrating and non-migrating cells. Cell Motil. Cytoskeleton 51, 27–38 (2002).

    Article  CAS  Google Scholar 

  18. Connelly, J. T. et al. Actin and serum response factor transduce physical cues from the microenvironment to regulate epidermal stem cell fate decisions. Nat. Cell Biol. 12, 711–718 (2010).

    Article  CAS  Google Scholar 

  19. Symons, M. H. & Mitchison, T. J. Control of actin polymerization in live and permeabilized fibroblasts. J. Cell Biol. 114, 503–513 (1991).

    Article  CAS  Google Scholar 

  20. Pollard, T. D., Blanchoin, L. & Mullins, R. D. Actin dynamics. J. Cell Sci. 114, 3–4 (2001).

    CAS  Google Scholar 

  21. Peng, G. E., Wilson, S. R. & Weiner, O. D. A pharmacological cocktail for arresting actin dynamics in living cells. Mol. Biol. Cell 22, 3986–3994 (2011).

    Article  CAS  Google Scholar 

  22. Mullins, R. D., Heuser, J. A. & Pollard, T. D. The interaction of Arp2/3 complex with actin: nucleation, high affinity pointed end capping, and formation of branching networks of filaments. Proc. Natl Acad. Sci. USA 95, 6181–6186 (1998).

    Article  CAS  Google Scholar 

  23. Yang, C. & Svitkina, T. Visualizing branched actin filaments in lamellipodia by electron tomography. Nat. Cell Biol. 13, 1012–1013 (2011).

    Article  CAS  Google Scholar 

  24. Sambeth, R. & Baumgaertner, A. Autocatalytic polymerization generates persistent random walk of crawling cells. Phys. Rev. Lett. 86, 5196–5199 (2001).

    Article  CAS  Google Scholar 

  25. Carlsson, A. E. Dendritic actin filament nucleation causes traveling waves and patches. Phys. Rev. Lett. 104, 228102 (2010).

    Article  Google Scholar 

  26. Burke, T. A. et al. Homeostatic actin cytoskeleton networks are regulated by assembly factor competition for monomers. Curr. Biol. 24, 579–585 (2014).

    Article  CAS  Google Scholar 

  27. Edelstein-Keshet, L. Mathematical Models in Biology (SIAM, 2005).

    Book  Google Scholar 

  28. Ruprecht, V. et al. Cortical contractility triggers a stochastic switch to fast amoeboid cell motility. Cell 160, 673–685 (2015).

    Article  CAS  Google Scholar 

  29. Shutova, M., Yang, C., Vasiliev, J. M. & Svitkina, T. Functions of nonmuscle myosin II in assembly of the cellular contractile system. PLoS ONE 7, e40814 (2012).

    Article  CAS  Google Scholar 

  30. Henson, J. H. et al. Two components of actin-based retrograde flow in sea urchin coelomocytes. Mol. Biol. Cell 10, 4075–4090 (1999).

    Article  CAS  Google Scholar 

  31. Ponti, A., Machacek, M., Gupton, S. L., Waterman-Storer, C. M. & Danuser, G. Two distinct actin networks drive the protrusion of migrating cells. Science 305, 1782–1786 (2004).

    Article  CAS  Google Scholar 

  32. Hu, K., Ji, L., Applegate, K. T., Danuser, G. & Waterman-Storer, C. M. Differential transmission of actin motion within focal adhesions. Science 315, 111–115 (2007).

    Article  CAS  Google Scholar 

  33. Mayer, M., Depken, M., Bois, J. S., Jülicher, F. & Grill, S. W. Anisotropies in cortical tension reveal the physical basis of polarizing cortical flows. Nature 467, 617–621 (2010).

    Article  CAS  Google Scholar 

  34. Koestler, S. A. et al. Arp2/3 complex is essential for actin network treadmilling as well as for targeting of capping protein and cofilin. Mol. Biol. Cell 24, 2861–2875 (2012).

    Article  Google Scholar 

  35. Henson, J. H. et al. Arp2/3 complex inhibition radically alters lamellipodial actin architecture, suspended cell shape, and the cell spreading process. Mol. Biol. Cell 26, 887–900 (2015).

    Article  CAS  Google Scholar 

  36. Kupfer, A., Louvard, D. & Singer, S. J. Polarization of the Golgi apparatus and the microtubule-organizing center in cultured fibroblasts at the edge of an experimental wound. Proc. Natl Acad. Sci. USA 79, 2603–2607 (1982).

    Article  CAS  Google Scholar 

  37. Bryant, D. M. & Mostov, K. E. From cells to organs: building polarized tissue. Nat. Rev. Mol. Cell Biol. 9, 887–901 (2008).

    Article  CAS  Google Scholar 

  38. Gause, G. F. The Struggle for Existence (Williams & Wilkins, 1934).

    Book  Google Scholar 

  39. Suarez, C. et al. Profilin regulates F-actin network homeostasis by favoring formin over Arp2/3 complex. Dev. Cell 32, 43–53 (2015).

    Article  CAS  Google Scholar 

  40. Rotty, J. D. et al. Profilin-1 serves as a gatekeeper for actin assembly by Arp2/3-dependent and -independent pathways. Dev. Cell 32, 54–67 (2015).

    Article  CAS  Google Scholar 

  41. Engl, W., Arasi, B., Yap, L. L., Thiery, J. P. & Viasnoff, V. Actin dynamics modulate mechanosensitive immobilization of E-cadherin at adherens junctions. Nat. Cell Biol. 16, 587–594 (2014).

    Article  CAS  Google Scholar 

  42. Wu, S. K. et al. Cortical F-actin stabilization generates apical-lateral patterns of junctional contractility that integrate cells into epithelia. Nat. Cell Biol. 16, 167–178 (2014).

    Article  CAS  Google Scholar 

  43. Wiggan, O., Shaw, A. E., DeLuca, J. G. & Bamburg, J. R. ADF/cofilin regulates actomyosin assembly through competitive inhibition of myosin II binding to F-actin. Dev. Cell 22, 530–543 (2012).

    Article  CAS  Google Scholar 

  44. Elam, W. A., Kang, H. & De La Cruz, E. M. Competitive displacement of cofilin can promote actin filament severing. Biochem. Biophys. Res. Commun. 438, 728–731 (2013).

    Article  CAS  Google Scholar 

  45. Hayakawa, K., Tatsumi, H. & Sokabe, M. Actin filaments function as a tension sensor by tension-dependent binding of cofilin to the filament. J. Cell Biol. 195, 721–727 (2011).

    Article  CAS  Google Scholar 

  46. McGrath, J. L., Osborn, E. A., Tardy, Y. S., Dewey, C. F. Jr & Hartwig, J. H. Regulation of the actin cycle in vivo by actin filament severing. Proc. Natl Acad. Sci. USA 97, 6532–6537 (2000).

    Article  CAS  Google Scholar 

  47. Omelchenko, T., Vasiliev, J. M., Gelfand, I. M., Feder, H. H. & Bonder, E. M. Mechanisms of polarization of the shape of fibroblasts and epitheliocytes: separation of the roles of microtubules and Rho-dependent actin-myosin contractility. Proc. Natl Acad. Sci. USA 99, 10452–10457 (2002).

    Article  CAS  Google Scholar 

  48. Omelchenko, T., Vasiliev, J. M., Gelfand, I. M., Feder, H. H. & Bonder, E. M. Rho-dependent formation of epithelial “leader” cells during wound healing. Proc. Natl Acad. Sci. USA 100, 10788–10793 (2003).

    Article  CAS  Google Scholar 

  49. Wang, H. R. et al. Regulation of cell polarity and protrusion formation by targeting RhoA for degradation. Science 302, 1775–1779 (2003).

    Article  CAS  Google Scholar 

  50. Asokan, S. B. et al. Mesenchymal chemotaxis requires selective inactivation of myosin II at the leading edge via a noncanonical PLCγ/PKCα pathway. Dev. Cell 31, 747–760 (2014).

    Article  CAS  Google Scholar 

  51. Lou, S. S., Diz-Muñoz, A., Weiner, O. D., Fletcher, D. A. & Theriot, J. A. Myosin light chain kinase regulates cell polarization independently of membrane tension or Rho kinase. J. Cell Biol. 209, 275–288 (2015).

    Article  CAS  Google Scholar 

  52. Lämmermann, T. & Sixt, M. Mechanical modes of ‘amoeboid’ cell migration. Curr. Opin. Cell Biol. 21, 636–644 (2009).

    Article  Google Scholar 

  53. Friedl, P. & Alexander, S. Cancer invasion and the microenvironment: plasticity and reciprocity. Cell 147, 992–1009 (2011).

    Article  CAS  Google Scholar 

  54. Maiuri, P. et al. Actin flows mediate a universal coupling between cell speed and cell persistence. Cell 161, 374–386 (2015).

    Article  CAS  Google Scholar 

  55. Ishizaki, T. et al. p160ROCK, a Rho-associated coiled-coil forming protein kinase, works downstream of Rho and induces focal adhesions. FEBS Lett. 404, 118–124 (1997).

    Article  CAS  Google Scholar 

  56. Pontrello, C. G. et al. Cofilin under control of β-arrestin-2 in NMDA-dependent dendritic spine plasticity, long-term depression (LTD), and learning. Proc. Natl Acad. Sci. USA 109, E442–E451 (2012).

    Article  CAS  Google Scholar 

  57. Nakayama, M. et al. Rho-kinase phosphorylates PAR-3 and disrupts PAR complex formation. Dev. Cell 14, 205–215 (2008).

    Article  CAS  Google Scholar 

  58. Kraynov, V. S. et al. Localized Rac activation dynamics visualized in living cells. Science 290, 333–337 (2000).

    Article  CAS  Google Scholar 

  59. Nalbant, P., Hodgson, L., Kraynov, V., Toutchkine, A. & Hahn, K. M. Activation of endogenous Cdc42 visualized in living cells. Science 305, 1615–1619 (2004).

    Article  CAS  Google Scholar 

  60. Subauste, M. C. et al. Rho family proteins modulate rapid apoptosis induced by cytotoxic T lymphocytes and Fas. J. Biol. Chem. 275, 9725–9733 (2000).

    Article  CAS  Google Scholar 

  61. Hao, J. J. et al. Phospholipase C-mediated hydrolysis of PIP2 releases ERM proteins from lymphocyte membrane. J. Cell Biol. 184, 451–462 (2009).

    Article  CAS  Google Scholar 

  62. Beach, J. R., Licate, L. S., Crish, J. F. & Egelhoff, T. T. Analysis of the role of Ser1/Ser2/Thr9 phosphorylation on myosin II assembly and function in live cells. BMC Cell Biol. 12, 52 (2011).

    Article  CAS  Google Scholar 

  63. Machacek, M. & Danuser, G. Morphodynamic profiling of protrusion phenotypes. Biophys. J. 90, 1439–1452 (2006).

    Article  CAS  Google Scholar 

  64. Ji, L. & Danuser, G. Tracking quasi-stationary flow of weak fluorescent signals by adaptive multi-frame correlation. J. Microsc. 220, 150–167 (2005).

    Article  CAS  Google Scholar 

  65. Mendoza, M. C., Besson, S. & Danuser, G. Quantitative fluorescent speckle microscopy (qFSM) to measure actin dynamics. Curr. Protoc. Cytom. 2 (2012).

Download references

Acknowledgements

The authors wish to acknowledge E. Bonder (Rutgers University, Newark, New Jersey, USA), T. Omelchenko (Sloan-Kettering Institute, New York, New York, USA) and J. Vasiliev (National Cancer Research Center, Moscow, Russia) for providing epithelial cell lines. We are grateful to I. Ethell (University of California Riverside, California, USA), S. Narumiya (Kyoto University, Japan), K. Kaibuchi (Nagoya University, Japan) and R. Horwitz (University of Virginia, Charlottesville, Virginia, USA) for sharing genetic constructs. We also thank M. Piel (Institut Curie, Paris, France) and D. Bonazzi (Institut Pasteur, Paris, France) for comments on the manuscript. This research was supported by a postdoctoral fellowship from the Leukemia & Lymphoma Society (grant no. 5388-13) to A.J.L., and the National Institutes of Health grants R01 GM071868 to G.D. and GM068952 to A.M. All light microscopy experiments described in the present work were performed at the Nikon Imaging Center of Harvard Medical School, Boston, Massachusetts.

Author information

Authors and Affiliations

Authors

Contributions

A.J.L., K.-C.L., A.M. and G.D. designed the project. A.J.L. and K.-C.L. performed all key experiments and analysed the data. D.A.B. performed 3D experiments. S.J.H. developed the software for membrane protrusivity and myosin flow analyses. M.D. generated fluorescence reporters for live-cell imaging experiments. A.J.L., A.M. and G.D. wrote the manuscript. All authors discussed the results and implications, and commented on the manuscript at all stages.

Corresponding authors

Correspondence to Alex Mogilner or Gaudenz Danuser.

Ethics declarations

Competing interests

The authors declare no competing financial interests.

Integrated supplementary information

Supplementary Figure 1 Quantitative analyses of large-scale morphodynamics in cultured epithelial cells IAR-2 in different signaling states.

(a) Cell shape descriptors (circularity and aspect ratio) and representative ‘masks’ of cells in a non-polarized and polarized state. (b) Shape descriptors measured in cells over-expressing wild-type (wt), constitutively active mutant (Q61L/Q63L; ‘ON-state’), and dominant negative mutant (T17N/T19N; ‘OFF-state’) versions of Cdc42, Rac1, RhoA, and the key components of the ROCK signaling pathway carrying activating (D) or inactivating (A) phosphomutations in ROCK-specific phosphorylation sites (n = 80 cells per each experimental condition; Mean ± SEM; P < 0.001, two-tailed unpaired Student’s t-test). (c) Modulation of RhoA-regulated formin-dependent actin polymerization, the microtubule cytoskeleton, vesicular trafficking, and chemotactic phosphoinositide signaling reactions per se is insufficient to induce migratory cell polarization in stationary epithelial cells. However, inhibiting RhoA/ROCK-dependent myosin-II activity alone produces cells with a polarized phenotype. Shape descriptors measured after 5 h of incubation with various small-molecule inhibitors (n = 120 cells per each experimental condition; Mean ± SEM; P < 0.001, two-tailed unpaired Student’s t-test). Targets of the inhibitors are specified in parentheses. (d) Proportion of cells remaining stationary, changing shape, or initiating motility within 4 h after addition of various inhibitors. Motility initiation was defined as an asymmetric, front-rear morphology and cell body displacement persisted over at least three cell lengths. Shape changers were non-polarized cells frequently switching between irregular shapes (n = 50 cells per each experimental condition; P < 0.001, two-tailed unpaired Student’s t-test).

Supplementary Figure 2 Increasing myosin-II activity switches migratory polarized epithelial cells back to stationary unpolarized morphologies.

Low levels of myosin-II activity in epithelial cells treated with 25 μm blebbistatin (BBS) are associated with spontaneous polarization and migration (Fig. 1). To increase myosin-II activity in such cells, BBS washout experiments were performed. The representative video sequence shows cells immediately after BBS washout. Scale bar, 10 μm. Removing BBS from the culture medium and thus increasing myosin-II activity restores typical stationary organization of epithelial cells, which is reflected in increased circularity, decreased aspect ratio and cell velocity (n = 30 cells, Mean ± SD—applies to all 3 graphs; P < 0.001, paired samples t-test).

Supplementary Figure 3 Isoform-specific effect of myosin-II on migratory cell polarity in epithelial cells.

(a) IAR-2 cells were transfected separately with nontargeting siRNA (Control), siRNA targeting Myh9 mRNA that encodes myosin-II A isoform, and siRNA targeting Myh10 mRNA that encodes myosin-II B isoform. After transfection, two types of cell mixtures (1:1) were obtained: (1) Myh9 siRNA + Control siRNA and (2) Myh10 siRNA + Control siRNA. Correlation of cell morphology (F-actin staining, red) with endogenous protein levels (myosin-II A- and myosin-II B-isoform-specific antibody staining, green) was conducted 72 h after transfection, as at this time minimal protein levels were detected. Unlike Myh10 siRNA-transfected cells, Myh9 siRNA-transfected epithelial cells lose actin rings, break cell-cell adhesions, and change shapes. Asterisk, Myh9 or Myh10 siRNA-transfected cells. Scale bar, 10 μm. (b) Western blotting-based evaluation of endogenous protein levels in cell lysates obtained 72 h after transfection with siRNA reagents. (c) Phase-contrast micrographs of live Myh9 or Myh10 siRNA-transfected cells. Next to each micrograph is a kymograph showing whole cell dynamics over time. Scale bar, 10 μm. (d) Phenotypic characteristics of cells treated with different siRNA reagents (n = 40 cells (Ctrl), 50 cells (Myh9), and 47 cells (Myh10); p(CtrlversusMyh9) < 0.001, p(CtrlversusMyh10) > 0.5, p(Myh9versusMyh10) < 0.001, two-tailed unpaired Student’s t-test). (e) Phenotypic characteristics of cells expressing different plasmid constructs and simultaneously treated with control or Myh9 siRNA reagents (n = 44 cells (siCtrl + Empty vector), 48 cells (siMyh9 + Empty vector), 45 cells (siMyh9 + wild-type [wt] myosin-IIA), and 48 cells (siMyh9 + myosin-IIAN93K27);p(siCtrl+EmptyvectorversussiMyh9+Emptyvector) < 0.001, p(siMyh9+EmptyvectorversussiMyh9+wtmyosin−IIA) < 0.001, p false( s i M y h 9  + wt myosin-IIA  v s . s i M y h 9  + myosin-IIA N 93 K 27 ) < 0.001, two-tailed unpaired Student’s t-test. Only cells expressing levels of exogenous protein comparable to that of endogenous protein were used for the analyses.

Supplementary Figure 4 Mathematical model predictions.

(a) Motility phase diagram and predicted cell shapes with actin network densities (blue, bundled actin; red, branched actin) below each shape. Thinner contours, earlier time points; thicker contours, later time points in stochastic simulations for Cell E (very high protrusion strength: a12 = 0.1, a21 > 3) and F (very high myosin strength: a12 > 3.0a21 = 0.1). (b) Dependence of the phase diagram on initial perturbations. Each simulation begins with a high bundled actin density everywhere. After some equilibration time (10 time units), a segment of bundled actin is ablated locally. The resulting bundled actin densities (blue) are step-function-like, with a sharp drop in the ablated area. The branched actin densities (red) are set to a low constant value. The width of the ablated bundled actin increases from left to right (left = 10%, middle = 20%, right = 50%). Top row, cases in which the drop in the bundled actin density is 80% of the initial value. Bottom row, cases in which the drop in the bundled actin density is 100% of the initial value. (c) Cell shape changes with increasing myosin inhibition. (d) Simulation of cells with stochastic F-actin dynamics.τ1 < τ2 < τ3 < τ4 < τ5, simulation time points. (e) 1D model with the global slow feedback. Top, simulation of the stochastic model without the feedback. The cell initially polarizes by establishing asymmetries in the density of branched versus bundled actin along the front-back axis. However, with time, stochastic fluctuations in myosin/actin densities can depolarize the cell. A representative case where bundled actin spontaneously ‘wins’ at the front is shown. Bottom, simulation of the stochastic model with the global feedback from long-range actin-myosin transport. The cell polarizes and is able to maintain its polarity because rearward actin-myosin transport and forward cell motion keep myosin density biased, dampening the stochastic fluctuation effect at the back. Red, branched actin at the front; Blue, bundled actin at the back; Magenta, branched actin at the back; Green, bundled actin at the front. X-axis, time; Y-axis, actin density (a.u.). Cartoon next to the graphs, a 2D interpretation of the simulations.

Supplementary Figure 5 Presence of serum in the growth medium is required for migratory cell polarization upon partial inactivation of myoin-II with blebbistatin.

IAR-2 cells were kept in fetal bovine serum (FBS)-free growth media for 5 h; after this step, the growth medium was changed and cells were cultivated for 3 h in the media containing 25 μM blebbistatin and 10% FBS (Serum add), or 25 μM blebbistatin and no FBS (No serum). By the end of the experiment, cells were fixed and stained for F-actin and nuclear DNA, and cell shape descriptors were statistically evaluated per each experimental condition (n = 80 cells for each case, Mean ± SEM; P < 0.001, two-tailed unpaired Student’s t-test). Scale bar, 10 μm.

Supplementary Figure 6 Motility-arrested cells fail to stabilize front-back asymmetry emerging upon myosin-II suppression.

The table displays percentage of cells with a specific morphodynamic behavior observed on substrates with uniform fibronectin coating (n = 45 cells) or on the microfabricated fibronectin pattern (n = 50 cells; p < 0.001, two-tailed unpaired Student’s t-test). The graph in which cell shape circularity index is plotted over time shows that BBS-treated cells on unpatterned fibronectin establish robust steady-state front-back asymmetry (low circularity, blue). In contrast, BBS-treated cells on circular fibronectin patterns rest in an unpolarized state (high circularity, red) or exhibit short bursts of frustrated front-back asymmetry that immediately return to an unpolarized state because of the missing ability to move forward (high circularity, green).

Supplementary Figure 7 Spatial differences in F-actin contractility exposed by localized cortical laser ablation (COLA) of the actin cytoskeleton in a polarized epithelial cell.

A two micron-sized laser cut was made at the branched actin-rich front and bundled actin-containing back pole of IAR-2 epithelial cells expressing mCherry-LifeAct and pretreated with BBS for 5 h (n = 10 cells; Mean ± SEM). The degree of local F-actin contractility at the front versus back of the cell was estimated from the initial recoil velocity of F-actin material adjacent to the laser cut. Assuming that contractility is a function of density of mechanochemically-active myosin-II motors and that the coefficient of friction is the same for actin filaments at the front and back poles of the cell, the analysis reveals a functional asymmetry of distinct cell poles in a migratory polarized cell: unlike the front, the back end of the cell displays contractile properties.

Supplementary Figure 8 Spatiotemporal organization of F-actin in Arp2/3-inhibited epithelial cells at low contractility regimes.

(a) F-actin organization in a myosin-inhibited cell with suppressed Arp2/3 activity (BBS + CK666). Scale bar, 5 μm. (b) F-actin dynamics in myosin-inhibited cells with functional Arp2/3 (BBS + CK689) and suppressed Arp2/3 activity (BBS + CK666). Zoomed fragments of cells are shown. F-actin (labeled with mCherry-LifeAct) was imaged with a 5-second interval for 60 s. Each timeframe from the movie was color-coded and the entire movie presented as a maximum projection of all timeframes. Color scale, time in seconds. Linescans, F-actin fluorescence intensity from the cell edge to the cell center. Scale bar, 1 μm.

Supplementary information

Supplementary Information

Supplementary Information (PDF 1661 kb)

A single IAR-2 epithelial cell 24 h after plating on a glass surface.

Differential interference contrast (DIC) videomicroscopy. Time, hours:minutes. Scale bar, 10 μm. (MOV 1475 kb)

Spontaneous symmetry breaking upon acute inhibition of myosin-II.

DIC movie is shown on the left; tracking of the cell boundary based on the time sequence of DIC images is shown on the right. IAR-2 cells were pretreated with 25 μM blebbistatin for 5 min and subsequently imaged in blebbistatin-containing medium. Time, hours:minutes. Scale bar, 10 μm. (MOV 760 kb)

A myosin-II-inhibited epithelial cell IAR-2 3 h after addition of the myosin-II drug blebbistatin.

DIC videomicroscopy. Time, hours:minutes. Scale bar, 10 μm. (MOV 1691 kb)

The recoil response of a marginal actin bundle upon localized cortical laser ablation (COLA).

A peripheral region of an IAR-2 cell expressing mEmerald-LifeAct and filmed by spinning disk confocal microscopy. The site of ablation is marked by a square; white arrows show the free ends of the ring upon ablation. Time, minutes:seconds. Scale bar, 3 μm. (MOV 459 kb)

Mass-balance between branched and bundled actin in cell shape and polarity determination (1).

At myosin strengths lower than in control situations, the bundled actomyosin and branched actin networks spatially segregate around the cell boundary while the cell polarizes, and stable cell motility ensues. The dynamic cell boundary is shown in the coordinate system moving with the geometric center of the cell; hence, the cell appears stationary, while in fact it is moving steadily to the right, which is shown below and also by the grid moving with the lab frame. Color scale, red—protrusion; blue—retraction. (MOV 349 kb)

Mass-balance between branched and bundled actin in cell shape and polarity determination (2).

In a control situation, the bundled actomyosin network effectively inhibits the branched actin networks all around the cell periphery. The cell remains symmetric and non-motile. Initial perturbations relax and do not destabilize the symmetric cell state. An intense perturbation is introduced closer to the end of the movie that wipes out part of the actomyosin bundle. The cell becomes motile temporarily and quickly reverts into the stationary symmetric circular state. Color scale, red—protrusion; blue—retraction. (MOV 76 kb)

Mass-balance between branched and bundled actin in cell shape and polarity determination (3).

At very low myosin strength, the branched actin network effectively inhibits the bundled actomyosin all around the periphery causing incoherent symmetry breaking of the cell but not coherent motility. Color scale, red—protrusion; blue—retraction. (MOV 114 kb)

Acute inhibition of myosin-II in stationary epithelial cells stimulates retrograde flow of myosin-II concomitant with the depletion of cortical myosin-II from the cell edge.

IAR-2 cells expressing mCherry-MRLC were pretreated with 25 μM blebbistatin for 5 min and subsequently imaged by spinning disk confocal microscopy in blebbistatin-containing medium. Time, minutes:seconds. Scale bar, 1 μm. (MOV 11403 kb)

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Lomakin, A., Lee, KC., Han, S. et al. Competition for actin between two distinct F-actin networks defines a bistable switch for cell polarization. Nat Cell Biol 17, 1435–1445 (2015). https://doi.org/10.1038/ncb3246

Download citation

  • Received:

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/ncb3246

This article is cited by

Search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing