Cellular chirality arising from the self-organization of the actin cytoskeleton

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Abstract

Cellular mechanisms underlying the development of left–right asymmetry in tissues and embryos remain obscure. Here, the development of a chiral pattern of actomyosin was revealed by studying actin cytoskeleton self-organization in cells with isotropic circular shape. A radially symmetrical system of actin bundles consisting of α-actinin-enriched radial fibres (RFs) and myosin-IIA-enriched transverse fibres (TFs) evolved spontaneously into the chiral system as a result of the unidirectional tilting of all RFs, which was accompanied by a tangential shift in the retrograde movement of TFs. We showed that myosin-IIA-dependent contractile stresses within TFs drive their movement along RFs, which grow centripetally in a formin-dependent fashion. The handedness of the chiral pattern was shown to be regulated by α-actinin-1. Computational modelling demonstrated that the dynamics of the RF–TF system can explain the pattern transition from radial to chiral. Thus, actin cytoskeleton self-organization provides built-in machinery that potentially allows cells to develop left–right asymmetry.

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Figure 1: Distinct patterns of actin cytoskeleton self-organization.
Figure 2: Composition and structure of RFs and TFs.
Figure 3: Probing of actin fibres in permeabilized cells with myosin-V-coated nanoparticles.
Figure 4: Dynamics of RFs and TFs.
Figure 5: The influence of Arp2/3-mediated actin polymerization, actomyosin contractility and microtubule integrity on the evolution of actin organization.
Figure 6: The effect of α-actinin-1 overexpression on the frequency and handedness of the chiral swirling.
Figure 7: The effects of α-actinin-1 dominant-negative mutant and knockdown.
Figure 8: Physical model for the actin self-organization.

References

  1. 1

    Blum, M., Feistel, K., Thumberger, T. & Schweickert, A. The evolution and conservation of left–right patterning mechanisms. Development 141, 1603–1613 (2014).

  2. 2

    Coutelis, J. B., Gonzalez-Morales, N., Geminard, C. & Noselli, S. Diversity and convergence in the mechanisms establishing L/R asymmetry in metazoa. EMBO Rep. 15, 926–937 (2014).

  3. 3

    Yoshiba, S. & Hamada, H. Roles of cilia, fluid flow, and Ca2+ signaling in breaking of left–right symmetry. Trends Genet. 30, 10–17 (2014).

  4. 4

    Wan, L. Q. & Vunjak-Novakovic, G. Micropatterning chiral morphogenesis. Commun. Integr. Biol. 4, 745–748 (2011).

  5. 5

    Wan, L. Q. et al. Micropatterned mammalian cells exhibit phenotype-specific left–right asymmetry. Proc. Natl Acad. Sci. USA 108, 12295–12300 (2011).

  6. 6

    Chen, T. H. et al. Left–right symmetry breaking in tissue morphogenesis via cytoskeletal mechanics. Circ. Res. 110, 551–559 (2012).

  7. 7

    Xu, J. et al. Polarity reveals intrinsic cell chirality. Proc. Natl Acad. Sci. USA 104, 9296–9300 (2007).

  8. 8

    Heacock, A. M. & Agranoff, B. W. Clockwise growth of neurites from retinal explants. Science 198, 64–66 (1977).

  9. 9

    Tamada, A., Kawase, S., Murakami, F. & Kamiguchi, H. Autonomous right-screw rotation of growth cone filopodia drives neurite turning. J. Cell Biol. 188, 429–441 (2010).

  10. 10

    Hagmann, J. Pattern formation and handedness in the cytoskeleton of human platelets. Proc. Natl Acad. Sci. USA 90, 3280–3283 (1993).

  11. 11

    Yamanaka, H. & Kondo, S. Rotating pigment cells exhibit an intrinsic chirality. Genes Cells 20, 29–35 (2015).

  12. 12

    Henley, C. L. Possible origins of macroscopic left–right asymmetry in organisms. J. Stat. Phys. 148, 741–775 (2012).

  13. 13

    Vandenberg, L. N., Lemire, J. M. & Levin, M. It’s never too early to get it Right: a conserved role for the cytoskeleton in left–right asymmetry. Commun. Integr. Biol. 6, e27155 (2013).

  14. 14

    Vasiliev, J. M. Spreading of non-transformed and transformed cells. Biochim. Biophys. Acta 780, 21–65 (1985).

  15. 15

    Small, J. V., Rottner, K., Kaverina, I. & Anderson, K. I. Assembling an actin cytoskeleton for cell attachment and movement. Biochim. Biophys. Acta 1404, 271–281 (1998).

  16. 16

    Tojkander, S., Gateva, G. & Lappalainen, P. Actin stress fibers–assembly, dynamics and biological roles. J. Cell Sci. 125, 1855–1864 (2012).

  17. 17

    Burridge, K. & Wittchen, E. S. The tension mounts: stress fibers as force-generating mechanotransducers. J. Cell Biol. 200, 9–19 (2013).

  18. 18

    Heath, J. P. Direct evidence for microfilament-mediated capping of surface receptors on crawling fibroblasts. Nature 302, 532–534 (1983).

  19. 19

    Hotulainen, P. & Lappalainen, P. Stress fibers are generated by two distinct actin assembly mechanisms in motile cells. J. Cell Biol. 173, 383–394 (2006).

  20. 20

    Naumanen, P., Lappalainen, P. & Hotulainen, P. Mechanisms of actin stress fibre assembly. J. Microsc. 231, 446–454 (2008).

  21. 21

    Vallenius, T. Actin stress fibre subtypes in mesenchymal-migrating cells. Open Biol. 3, 130001 (2013).

  22. 22

    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).

  23. 23

    Tojkander, S. et al. A molecular pathway for myosin II recruitment to stress fibers. Curr. Biol. 21, 539–550 (2011).

  24. 24

    Kovac, B., Teo, J. L., Makela, T. P. & Vallenius, T. Assembly of non-contractile dorsal stress fibers requires α-actinin-1 and Rac1 in migrating and spreading cells. J. Cell Sci. 126, 263–273 (2013).

  25. 25

    Gateva, G., Tojkander, S., Koho, S., Carpen, O. & Lappalainen, P. Palladin promotes assembly of non-contractile dorsal stress fibers through VASP recruitment. J. Cell Sci. 127, 1887–1898 (2014).

  26. 26

    Shemesh, T., Verkhovsky, A. B., Svitkina, T. M., Bershadsky, A. D. & Kozlov, M. M. Role of focal adhesions and mechanical stresses in the formation and progression of the lamellipodium-lamellum interface. Biophys. J. 97, 1254–1264 (2009); correction 97, 2115 (2009)

  27. 27

    Burnette, D. T. et al. A role for actin arcs in the leading-edge advance of migrating cells. Nat. Cell Biol. 13, 371–381 (2011).

  28. 28

    Burnette, D. T. et al. A contractile and counterbalancing adhesion system controls the 3D shape of crawling cells. J. Cell Biol. 205, 83–96 (2014).

  29. 29

    Sjöblom, B., Salmazo, A. & Djinović-Carugo, K. α-Actinin structure and regulation. Cell. Mol. Life Sci. 65, 2688–2701 (2008).

  30. 30

    Hariadi, R. F., Cale, M. & Sivaramakrishnan, S. Myosin lever arm directs collective motion on cellular actin network. Proc. Natl Acad. Sci. USA 111, 4091–4096 (2014).

  31. 31

    Mattila, P. K. & Lappalainen, P. Filopodia: molecular architecture and cellular functions. Nat. Rev. Mol. Cell Biol. 9, 446–454 (2008).

  32. 32

    Rizvi, S. A. et al. Identification and characterization of a small molecule inhibitor of formin-mediated actin assembly. Chem. Biol. 16, 1158–1168 (2009).

  33. 33

    Nolen, B. J. et al. Characterization of two classes of small molecule inhibitors of Arp2/3 complex. Nature 460, 1031–1034 (2009).

  34. 34

    Suraneni, P. et al. The Arp2/3 complex is required for lamellipodia extension and directional fibroblast cell migration. J. Cell Biol. 197, 239–251 (2012).

  35. 35

    Wu, C. et al. Arp2/3 is critical for lamellipodia and response to extracellular matrix cues but is dispensable for chemotaxis. Cell 148, 973–987 (2012).

  36. 36

    Straight, A. F. et al. Dissecting temporal and spatial control of cytokinesis with a myosin II inhibitor. Science 299, 1743–1747 (2003).

  37. 37

    Uehata, M. et al. Calcium sensitization of smooth muscle mediated by a Rho-associated protein kinase in hypertension. Nature 389, 990–994 (1997).

  38. 38

    Hoffman, L. M. et al. Genetic ablation of zyxin causes Mena/VASP mislocalization, increased motility, and deficits in actin remodeling. J. Cell Biol. 172, 771–782 (2006).

  39. 39

    Djinovic-Carugo, K., Young, P., Gautel, M. & Saraste, M. Structure of the α-actinin rod: molecular basis for cross-linking of actin filaments. Cell 98, 537–546 (1999).

  40. 40

    Choi, C. K. et al. Actin and α-actinin orchestrate the assembly and maturation of nascent adhesions in a myosin II motor-independent manner. Nat. Cell Biol. 10, 1039–1050 (2008).

  41. 41

    Roca-Cusachs, P. et al. Integrin-dependent force transmission to the extracellular matrix by α-actinin triggers adhesion maturation. Proc. Natl Acad. Sci. USA 110, E1361–E1370 (2013).

  42. 42

    Low, S. H., Mukhina, S., Srinivas, V., Ng, C. Z. & Murata-Hori, M. Domain analysis of α-actinin reveals new aspects of its association with F-actin during cytokinesis. Exp. Cell Res. 316, 1925–1934 (2010).

  43. 43

    Kozlov, M. M. & Bershadsky, A. D. Processive capping by formin suggests a force-driven mechanism of actin polymerization. J. Cell Biol. 167, 1011–1017 (2004).

  44. 44

    Shemesh, T. & Kozlov, M. M. Actin polymerization upon processive capping by formin: a model for slowing and acceleration. Biophys. J. 92, 1512–1521 (2007).

  45. 45

    Cramer, L. P., Siebert, M. & Mitchison, T. J. Identification of novel graded polarity actin filament bundles in locomoting heart fibroblasts: implications for the generation of motile force. J. Cell Biol. 136, 1287–1305 (1997).

  46. 46

    Paul, A. S. & Pollard, T. D. Review of the mechanism of processive actin filament elongation by formins. Cell Motil. Cytoskeleton 66, 606–617 (2009).

  47. 47

    Breitsprecher, D. & Goode, B. L. Formins at a glance. J. Cell Sci. 126, 1–7 (2013).

  48. 48

    Xu, Y. et al. Crystal structures of a Formin Homology-2 domain reveal a tethered dimer architecture. Cell 116, 711–723 (2004).

  49. 49

    Shemesh, T., Otomo, T., Rosen, M. K., Bershadsky, A. D. & Kozlov, M. M. A novel mechanism of actin filament processive capping by formin: solution of the rotation paradox. J. Cell Biol. 170, 889–893 (2005).

  50. 50

    Mizuno, H. et al. Rotational movement of the formin mDia1 along the double helical strand of an actin filament. Science 331, 80–83 (2011).

  51. 51

    Courtemanche, N., Lee, J. Y., Pollard, T. D. & Greene, E. C. Tension modulates actin filament polymerization mediated by formin and profilin. Proc. Natl Acad. Sci. USA 110, 9752–9757 (2013).

  52. 52

    Jegou, A., Carlier, M. F. & Romet-Lemonne, G. Formin mDia1 senses and generates mechanical forces on actin filaments. Nat. Commun. 4, 1883 (2013).

  53. 53

    Soranno, T. & Bell, E. Cytostructural dynamics of spreading and translocating cells. J. Cell Biol. 95, 127–136 (1982).

  54. 54

    Speder, P., Adam, G. & Noselli, S. Type ID unconventional myosin controls left–right asymmetry in Drosophila. Nature 440, 803–807 (2006).

  55. 55

    Hozumi, S. et al. An unconventional myosin in Drosophila reverses the default handedness in visceral organs. Nature 440, 798–802 (2006).

  56. 56

    Speder, P. & Noselli, S. Left–right asymmetry: class I myosins show the direction. Curr. Opin. Cell Biol. 19, 82–87 (2007).

  57. 57

    Naganathan, S. R., Furthauer, S., Nishikawa, M., Julicher, F. & Grill, S. W. Active torque generation by the actomyosin cell cortex drives left–right symmetry breaking. eLife 3, e04165 (2014).

  58. 58

    Fürthauer, S., Strempel, M., Grill, S. W. & Jülicher, F. Active chiral processes in thin films. Phys. Rev. Lett. 110, 048103 (2013).

  59. 59

    Thery, M., Pepin, A., Dressaire, E., Chen, Y. & Bornens, M. Cell distribution of stress fibres in response to the geometry of the adhesive environment. Cell Motil. Cytoskeleton 63, 341–355 (2006).

  60. 60

    Anderson, T. F. Techniques for the preservation of 3-dimensional structure in preparing specimens for the electron microscope. Trans. N. Y. Acad. Sci. 13, 130–134 (1951).

  61. 61

    Buckley, I. K. & Porter, K. R. Electron-microscopy of critical-point dried whole cultured-cells. J. Microsc. 104, 107–120 (1975).

  62. 62

    Mastronarde, D. N. Automated electron microscope tomography using robust prediction of specimen movements. J. Struct. Biol. 152, 36–51 (2005).

  63. 63

    Kremer, J. R., Mastronarde, D. N. & McIntosh, J. R. Computer visualization of three-dimensional image data using IMOD. J. Struct. Biol. 116, 71–76 (1996).

  64. 64

    Bryant, Z., Altman, D. & Spudich, J. A. The power stroke of myosin VI and the basis of reverse directionality. Proc. Natl Acad. Sci. USA 104, 772–777 (2007).

  65. 65

    Rock, R. S., Rief, M., Mehta, A. D. & Spudich, J. A. In vitro assays of processive myosin motors. Methods 22, 373–381 (2000).

  66. 66

    Rothemund, P. W. K. Folding DNA to create nanoscale shapes and patterns. Nature 440, 297–302 (2006).

  67. 67

    Sivaramakrishnan, S. & Spudich, J. A. Coupled myosin VI motors facilitate unidirectional movement on an F-actin network. J. Cell Biol. 187, 53–60 (2009).

  68. 68

    Tint, I. S., Hollenbeck, P. J., Verkhovsky, A. B., Surgucheva, I. G. & Bershadsky, A. D. Evidence that intermediate filament reorganization is induced by Atp-dependent contraction of the actomyosin cortex in permeabilized fibroblasts. J. Cell Sci. 98, 375–384 (1991).

  69. 69

    Kerssemakers, J. W. et al. Assembly dynamics of microtubules at molecular resolution. Nature 442, 709–712 (2006).

  70. 70

    Gelles, J., Schnapp, B. J. & Sheetz, M. P. Tracking kinesin-driven movements with nanometre-scale precision. Nature 331, 450–453 (1988).

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Acknowledgements

We thank P. Lappalainen for discussion, S. Hanks, B. M. Jockush, I. Kaverina, C. Otey, P. Roca-Cusachs, M. J. Schell and R. Wedlich-Soldner for providing reagents, C. Lu for writing the custom script for velocities measurement, Z. Z. Lieu for help in the knockdown study, S. Wolf for expert help in paper editing, the microscopy core facility at the Mechanobiology Institute for technical help and Sanford Burnham Medical Research Institute for electron microscopy work. This research has been supported by the National Research Foundation Singapore, Ministry of Education of Singapore, Grant R-714-006-006-271, and administrated by the National University of Singapore. K.L.A. and D.H. were supported by National Institutes of Health (NIH) grant P01-GM098412. C.P. and N.V. were supported by NIH grant P01-GM066311. M.M.K. was supported by the Israel Science Foundation (grant No.758/11) and the Marie Curie network Virus Entry, and holds the Joseph Klafter Chair in Biophysics. M.M.K. thanks the Mechanobiology Institute, National University of Singapore, for hospitality. A.D.B. holds the Joseph Moss Professorial Chair in Biomedical Research at the Weizmann Institute and is a Visiting Professor at the National University of Singapore and acknowledges support from the Israel Science Foundation (grant No. 956/10).

Author information

A.D.B. conceived the study. Y.H.T., T.S., M.M.K. and A.D.B. designed the study, analysed the data and wrote the manuscript with input from all authors. Y.H.T. and V.T. performed most experiments. R.F.H. and S.S. performed the cytoskeletal probing with myosin-V- and -VI-coated nanoparticles and analysed the data. K.L.A., C.P., N.V. and D.H. performed the electron microscopy work comprising of correlative light and electron microscopy, electron tomography and tomographic three-dimensional reconstructions. T.S. and M.M.K. developed the computational model.

Correspondence to Tom Shemesh or Alexander D. Bershadsky.

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The authors declare no competing financial interests.

Integrated supplementary information

Supplementary Figure 1 Overexpression of radial fiber components, Zyxin or VASP, is not sufficient to reverse chirality.

RFP-Zyxin (a) and GFP-VASP (c) localized to radial fibers and focal adhesions. Actin was visualized either by co-transfection with LifeAct-GFP (green in a) or phalloidin staining (red in c). Scale bars, 10 μm. RFP-Zyxin (b) or GFP-VASP (d) expressing cells were filmed to follow their actin dynamics. The expression level in an individual cell was assessed by measuring either total RFP or total GFP fluorescence intensity respectively. The y-axis represents fluorescence intensity in a log scale; the cells were sorted in ascending order of their respective protein expression level along the x-axis. Each bar on the histogram represents an individual cell. Colour-coding indicates the type of actin cytoskeleton dynamics for each cell. Green – cells that do not demonstrate swirling during the entire period of observation (10 hours). Blue – cells demonstrating positive tilt of radial fibers and counter-clockwise swirling similar to control cells. No clockwise swirling events were observed in these experiments. 58 cells assessed for RFP-Zyxin, and 72 cells assessed for GFP-VASP.

Supplementary Figure 2 Electron tomography showing 3D morphology and organization of transverse and radial fibers.

(ad) Surface representations of overlapping 3D electron tomographic reconstructions running along radial fibers. The relative spatial arrangement of the four reconstructions is shown below d. The center of the cell is towards the left of a, while the cell edge is shown in d. The height (in nm) of the features are mapped according to the colour key on the right of a. The area of a reconstructed image is 2.8 μm2. Radial (RF) and transverse (TF) fibers as well as the transversal layer (TL) are indicated in b. Nearer to the cell center (a,b), the transverse and radial fibers (blue and cyan) are above the transversal layer (orange). Towards the cell edge (c,d), the two fiber systems move closer to the substrate (colour shift from blue/cyan to cyan/green and green/orange) and finally appear below the orange transversal layer plane (in d) when the focal adhesions contact the substrate. (e) Cross-section of the reconstruction shown in a. The upper panel shows a 10 nm thick cross-section along the long axis of the radial fibers to the top of a, perpendicular to the transverse fibers, as indicated in the scheme at the upper left corner of e. The transverse layer (TL) and the substrate (S) locations are marked at the sides of the figure. The location of the radial fiber (RF) and the location where the transverse fiber (TF) runs perpendicular to the image plane are also marked. Transverse fibers appear to interact with radial fibers. The height (in nm) is shown in the bar on the right. The bar is colour-coded for comparison with ad. The width of the cross-section is 1.9 μm. The components of the radial fibers go in and out of the cross-section, generating an alternating pattern of black and white regions. The lower panel shows the model derived from the tomographic reconstructions in a view similar to that in the upper panel for comparison. (f) XY slices through the tomographic reconstruction (each 2 nm thick) shown in b (see also Supplementary Video 4). The heights of the sections are indicated by the colour of the rim, which correspond to the colour-coding in a and e. Orthogonal side views are also shown for each slice on the top and to the right. The location of the XZ slices (top) within the XY view are indicated by the end of the red lines at the right of the XY view; the location of the YZ slices (right) are indicated by the end of the green lines at the top of the XY view. Radial fibers (RF), transversal layer (TL), as well as the top and bottom occurrences of the transverse fibers (TF) are marked on the slices. See also Supplementary Video 3 for comparison with fluorescence microscopy.

Supplementary Figure 3 Probing actin fibers in permeabilized cells with myosin VI-coated nanoparticles.

(a) Movement trajectories (green) of fluorescent nanoparticles coated with myosin VI along the actin cytoskeleton (red) of a fixed extracted cell. The white box marks the view shown in b. Scale bar, 10 μm. (be) High magnification view of myosin VI nanoparticle movement along the single transverse fiber. (b) A view of actin cytoskeleton (red) after extraction. White-boxed region: The transverse fiber decorated with myosin VI nanoparticles (green) is marked by the dotted line. (ce) Sequential images from the live imaging of myosin VI nanoparticles moving along the transverse fiber. Nanoparticles #1 and #2 (white arrowheads) are moving away from each other. Scale bars, 1 μm. The full-length sequence is shown in Supplementary Video 6.

Supplementary Figure 4 Self-organization of the actin cytoskeleton in cells with depolymerized microtubules.

Disruption of microtubules by nocodazole was confirmed by anti-α-tubulin immunofluorescence staining. Actin was visualized with phalloidin staining. The radial and chiral patterns of the actin cytoskeleton organization in cells with an intact microtubule system and in cells lacking microtubules are shown. Scale bars, 10 μm. See also Supplementary Video 10.

Supplementary Figure 5 Long “single-ended” stress-fibers replace radial fibers in GFP-ABDdel-α-actinin-1 expressing cells.

(a) Schematic diagram illustrating the molecular organization of α-actinin-1 and its mutants. The α-actinin-1 monomer has an N-terminal actin-binding domain (ABD) composed of two calponin-homology (CH) domains, a central rod consisting of four spectrin repeats (R1-R4), and a C-terminal calmodulin (CaM)-like domain. α-actinin-1 monomers form an anti-parallel dimer with the ABD domain present at both ends. The ABDdel-α-actinin-1 mutant lacks the ABD domain41. Spectrin repeats 1 and 2 are responsible for integrin binding. The SR12 mutant consists only of these spectrin repeats (R1 and R2)41. (b) Cells expressing GFP-ABDdel-α-actinin-1 were extracted prior to fixation to reduce background due to the presence of excess soluble GFP-ABDdel-α-actinin-1. Truncated GFP-ABDdel-α-actinin-1 mutant protein localized to long actin fibers (purple). Unlike radial fibers, these fibers also contained myosin-IIA (blue). However, unlike ventral stress-fibers, they were associated with only one focal adhesion (labeled by vinculin, red). Thus, we classified this type of actin fibers as “single-ended” stress-fibers. Scale bar, 10 μm. (c) Histogram showing the distribution of “single-ended” actin fiber tilt angle (°) in a cell expressing GFP-ABDdel-α-actinin-1. 30 actin fibers of a representative cell were analyzed. See also Supplementary Video 12. Note that unlike radial fibers in cells overexpressing the full-length α-actinin-1, the “single-ended” stress-fibers in cells expressing GFP-ABDdel-α-actinin-1 tilt in the same direction as radial fibers in control cells (see Fig. 6c).

Supplementary Figure 6 The effect of SR12-GFP α-actinin-1 fragment on actin cytoskeleton self-organization.

(a,b) Actin organization (visualized by tdTomato-F-Tractin) in SR12-GFP expressing cells and high magnification time-series images illustrating actin dynamics (white-boxed region). 60% of SR12-GFP transfected cells demonstrated phenotypes similar to control cells (30% counter-clockwise swirling and 30% non-swirling). The remaining 40% of the transfected cells were found to express a higher level (2-fold on average) of SR12-GFP. 60% of these higher SR12-GFP expressing cells failed to progress beyond the circular stage (a) while the remaining 40% exhibited similar behavior to GFP-ABDdel-α-actinin-1 mutant expressing cells, forming characteristic “single-ended” stress-fibers that tilt in a positive direction (b). An example of two “single-ended” stress-fibers are shown here (white and cyan arrowheads). Scale bars, 10 μm; 5 μm (high-magnification). n = 39 transfected cells pooled from 2 independent experiments.

Supplementary Figure 7 Membrane-bound beads move in a chiral motion coupled to the swirling of the actin network.

Fibronectin-coated beads (4.5 μm in diameter) moved in a counter-clockwise direction on the cell surface as the actin cytoskeleton swirled in the same direction. Yellow, red and cyan asterisks () marks the bead position at time points 0-, 14- and 38-minute respectively. The arrow indicates the direction of the bead movement. Scale bar, 10 μm.

Supplementary Figure 8 Schematic illustration depicting polymerization of individual actin filaments nucleated by formins.

(a) Polarized actin filaments (green) capped by a formin dimer (blue-red). The first three actin subunits in the two actin helices are numbered 1–3 and 1’-3’ respectively. The formin dimer is immobilized while the opposite end of the actin filament is not. Unconstrained actin filaments rotate in the positive angular direction about x-axis (magenta arrow), as experimentally shown by Mizuno et al.50. (b) Actin filaments are prevented from rotating via immobilization of barbed end by formin and trapping of opposite end by α-actinin cross-linking. Immobile actin subunits are labeled in yellow. Torsional elastic energy accumulated during polymerization is periodically released via high amplitude rotation (“screw-step” highlighted by gray background) in the negative direction (yellow arrow). See details in Shemesh et al.49. See also Supplementary Videos 17 and 18).

Supplementary Figure 9 Primary immunoblotting data.

(a) 10 μg samples of protein cell lysates from control siRNA- and ACTN1 siRNA-transfected cells were separated on SDS-PAGE gel and transferred to PVDF membranes. Positions of molecular weight markers are indicated to the right of each PVDF membrane. (b) Immunoblotting was performed with corresponding antibodies, anti-ACTN1, anti-ACTN4 and anti-α-tubulin, and developed using standard ECL procedure. Scans of uncropped immunoblot is shown here. Data from this immunoblot was used in Fig. 7c.

Supplementary information

Supplementary Information

Supplementary Information (PDF 3114 kb)

A typical example of the self-organization of the actin cytoskeleton in cell on a circular adhesive island.

Images were recorded at 2 minute intervals over a period of 11 hours. Display rate is 30 frames/sec and corresponds to the time-lapse series in Fig. 1b. (MOV 26775 kb)

Evolution of the actin cytoskeleton from the radial into the chiral pattern.

Images were recorded at 2 minute intervals over a period of 4 hours. Display rate is 7 frames/sec and corresponds to the time-lapse series in Fig. 5 (untreated). (MOV 1094 kb)

Organization of radial and transverse fibers seen in confocal microscopy Z-section.

Cells were fixed and stained with phalloidin to visualize actin. Images of the actin structures at the cell edge of non-dehydrated cell on a circular island were captured by Z-stepping at 0.25 μm per step from the bottom to the top of the cell (MOV 217 kb)

Electron tomographic reconstruction of intersection between transverse and radial fibers.

Movie showing successive 2-nm slices of the tomographic reconstruction (10 frames/sec) as shown in Supplementary Fig. 2f. The image display starts from the top, goes to the bottom, and then back up to the top. (MOV 10393 kb)

Movement of myosin V- or VI-coated nanoparticles (green) along filopodial cell projections (red) of fixed extracted cell.

An asterisk () indicates a filopodium. Myosin V-coated nanoparticles moved towards filament barbed ends away from the cell body, while myosin VI-coated nanoparticles moved towards filament pointed ends in the direction of the cell body. Images were recorded at half a second intervals. Display rate is 30 frames/sec. (MOV 504 kb)

Movement of myosin V- (left) and myosin VI-coated (right) nanoparticles (green) along the actin transverse fiber (red) of fixed extracted cell.

Note that the movements of both myosin V- and VI-coated nanoparticles could proceed in opposite directions along the same transverse fibers. Images were recorded at half a second intervals. Display rate is 7 frames/sec and corresponds to Fig. 3d–f (myosin V) and Supplementary Fig. 3b–e (myosin VI). (MOV 648 kb)

Actin dynamics upon treatment with the 10 μM formin inhibitor, SMIFH2.

Images were recorded at 2 minute intervals over a period of 3.5 hours. Display rate is 20 frames/sec and corresponds to the time-lapse series in Fig. 4a (lower panel). (MOV 2554 kb)

Growth of radial fiber is dependent on formin activity.

SMIFH2-treated cell was monitored for 1 hour prior to drug washout. The recovery phase was followed for 14 hours with images taken at 2 minute interval. Display rate is 30 frames/sec. Note the recovery of radial fibers growth started at 3 hours after drug washout, at the cell periphery. (MOV 2230 kb)

Actin dynamics upon treatment with 100 μM Y27632 (left) or 50 μM blebbistatin (Bleb, right).

Images were recorded at 2 minute intervals over a period of 5 hours. Display rate is 20 frames/sec and corresponds to the time-lapse series in Fig. 5. (MOV 728 kb)

Actin dynamics upon treatment with 1 μM nocodazole.

Images were recorded at 2 minute intervals over a period of 5 hours. Note that depolymerization of microtubules does not interfere with formation of chiral pattern and does not change handedness of the actin swirl. Display rate is 20 frames/sec and corresponds to the time-lapse series in Fig. 5. (MOV 970 kb)

Handedness of the chiral pattern in cell expressing low (left) or high (right) level of GFP-α-actinin-1.

Radial fibers were visualized with GFP-α-actinin-1. Images were recorded at 2 minute intervals over a period of 6 hours. Display rate is 20 frames/sec and corresponds to Fig. 6a. (MOV 3581 kb)

A typical example of actin dynamics in cell expressing truncated mutant GFP-ABDdel-α-actinin-1.

Images were recorded at 4 minute intervals over a period of 9 hours. Display rate is 6 frames/sec and corresponds to the time-lapse series in Fig. 7a. (MOV 4573 kb)

A typical example of actin dynamics in α-actinin-1 knockdown cell.

Images were recorded at 3 minute intervals over a period of 9 hours. Display rate is 8 frames/sec and corresponds to the time-lapse series in Fig. 7f. (MOV 3954 kb)

Rotation of the microtubule array in the counter-clockwise direction, together with counter-clockwise swirling of the actin network, seen in cell expressing GFP-tubulin and tdTomato-F-Tractin to visualize microtubule and actin respectively.

Images were recorded at 2 minute intervals over a period of 12 hours. Display rate is 30 frames/sec. (MOV 3407 kb)

Simulation of the chiral self-organization of the actin cytoskeleton.

Corresponds to Fig. 8b. (MOV 717 kb)

Simulation showing polymerization of a free actin filament capped by formin-FH2 dimer.

Actin monomers are shown in green (dark and light green are used to distinguish the long pitch helices). FH2 dimer is shown in blue-red. Note the rotation of the formin dimer. (MOV 1308 kb)

Simulation showing polymerization of actin filament capped by an immobilized formin-FH2 dimer.

Actin monomers are shown in green (dark and light green are used to distinguish the long pitch helices). FH2 dimer is shown in blue-red. Note the rotation of the actin filament. (MOV 1540 kb)

Simulation showing polymerization of actin filament capped by an immobilized formin-FH2 dimer, and prevented from rotating at the pointed end.

Actin monomers are shown in green (dark and light green are used to distinguish the long pitch helices). FH2 dimer is shown in blue-red. Immobilized monomers at the tip of the filament are shown in yellow. Note the periodic high amplitude rotation of the actin filament upon transit release of the contact of its barbed end with formin dimer (“screw-steps”; highlighted by change in background colour from black to light gray). (MOV 1439 kb)

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Tee, Y., Shemesh, T., Thiagarajan, V. et al. Cellular chirality arising from the self-organization of the actin cytoskeleton. Nat Cell Biol 17, 445–457 (2015) doi:10.1038/ncb3137

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