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Lattice defects induce microtubule self-renewal

Nature Physics volume 15, pages 830–838 (2019)Cite this article

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Abstract

Microtubules are dynamic polymers, which grow and shrink by addition and removal of tubulin dimers at their extremities. Within the microtubule shaft, dimers adopt a densely packed and highly ordered crystal-like lattice structure, which is generally not considered to be dynamic. Here, we report that thermal forces are sufficient to remodel the microtubule shaft, despite its apparent stability. Our combined experimental data and numerical simulations on lattice dynamics and structure suggest that dimers can spontaneously leave and be incorporated into the lattice at structural defects. We propose a model mechanism, where the lattice dynamics is initiated via a passive breathing mechanism at dislocations, which are frequent in rapidly growing microtubules. These results show that we may need to extend the concept of dissipative dynamics, previously established for microtubule extremities, to the entire shaft, instead of considering it as a passive material.

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Fig. 1: Incorporation of free tubulin into the microtubule lattice visualized by TIRF.
Fig. 2: Monte Carlo simulations of microtubule lattice dynamics.
Fig. 3: Dislocation defects in the microtubule lattice detected by cryo-electron microscopy.
Fig. 4: Incorporation of free tubulin into the microtubule lattice visualized by TIRF.
Fig. 5

Data availability

The data that support the findings of this study are available from the authors upon reasonable request, see author contributions for specific datasets.

Code availability

The source code of the kinetic Monte Carlo model along with detailed instructions to reproduce the data for this manuscript is available online (https://sourceforge.net/projects/microtubulelatticemodel). The codes for the analysis of the model simulations are available from the corresponding authors upon request.

References

  1. Carlier, M.-F. Guanosine-5′-triphosphate hydrolysis and tubulin polymerization. Mol. Cell. Biochem. 47, 97–113 (1982).

    Article  Google Scholar 

  2. Mitchison, T. & Kirschner, M. W. Dynamic instability of microtubule growth. Nature 312, 237–242 (1984).

    Article  ADS  Google Scholar 

  3. Walker, R. A. et al. Dynamic instability of individual microtubules analyzed by video light microscopy: rate constants and transition frequencies. J. Cell Biol. 107, 1437–1448 (1988).

    Article  Google Scholar 

  4. Howard, J. & Hyman, A. A. Dynamics and mechanics of microtubule plus end. Nature 422, 753–758 (2003).

    Article  ADS  Google Scholar 

  5. Duellberg, C., Cade, N. I., Holmes, D. & Surrey, T. The size of the EB cap determines instantaneous microtubule stability. eLife 5, e13470 (2016).

    Article  Google Scholar 

  6. Aher, A. & Akhmanova, A. Tipping microtubule dynamics, one protofilament at a time. Curr. Opin. Cell Biol. 50, 86–92 (2018).

    Article  Google Scholar 

  7. Akhmanova, A. & Steinmetz, M. O. Control of microtubule organization and dynamics: two ends in the limelight. Nat. Rev. Mol. Cell Biol. 16, 711–726 (2015).

    Article  Google Scholar 

  8. Gasic, I. & Mitchison, T. J. Autoregulation and repair in microtubule homeostasis. Curr. Opin. Cell Biol. 56, 80–87 (2019).

    Article  Google Scholar 

  9. Dye, R. B., Flicker, P. F., Lien, D. Y. & Williams, R. C. End-stabilized microtubules observed in vitro: stability, subunit, interchange, and breakage. Cell Motil. Cytoskel. 21, 171–186 (1992).

    Article  Google Scholar 

  10. Schaedel, L. et al. Microtubules self-repair in response to mechanical stress. Nat. Mater. 14, 1156–1163 (2015).

    Article  ADS  Google Scholar 

  11. Reid, T. A., Coombes, C. & Gardner, M. K. Manipulation and quantification of microtubule lattice integrity. Biol. Open 6, 1245–1256 (2017).

    Article  Google Scholar 

  12. Dimitrov, A. et al. Detection of GTP-tubulin conformation in vivo reveals a role for GTP remnants in microtubule rescue. Science 322, 1353–1356 (2008).

    Article  ADS  Google Scholar 

  13. de Forges, H. et al. Localized mechanical stress promotes microtubule rescue. Curr. Biol. 26, 3399–3406 (2016).

    Article  Google Scholar 

  14. Aumeier, C. et al. Self-repair promotes microtubule rescue. Nat. Cell Biol. 18, 1054–1064 (2016).

    Article  Google Scholar 

  15. Vemu, A. et al. Severing enzymes amplify mirotubule arrays through lattice GTP-tubulin incorporation. Science 361, eaau1504 (2018).

    Article  Google Scholar 

  16. VanBuren, V., Odde, D. J. & Cassimeris, L. Estimations of lateral and longitudinal bond energies within the microtubule lattice. Proc. Natl Acad. Sci. USA 99, 6035–6040 (2002).

    Article  ADS  Google Scholar 

  17. VanBuren, V., Cassimeris, L. & Odde, D. J. Mechanochemical model of microtubule structure and self-assembly kinetics. Biophys. J. 89, 2911–2926 (2005).

    Article  Google Scholar 

  18. Sept, D., Baker, N. A. & McCammon, J. A. The physical basis of microtubule structure and stability. Prot. Sci. 12, 2257–2261 (2003).

    Article  Google Scholar 

  19. Chrétien, D. & Fuller, S. D. Microtubules switch occasionally into unfavorable configurations during elongation. J. Mol. Biol. 298, 663–676 (2000).

    Article  Google Scholar 

  20. Chrétien, D., Metoz, F., Verde, F., Karsenti, E. & Wade, R. H. Lattice defects in microtubules: protofilament numbers vary within individual microtubules. J. Cell Biol. 117, 1031–1040 (1992).

    Article  Google Scholar 

  21. Atherton, J., Stouffer, M., Francis, F. & Moores, C. A. Microtubule architecture in vitro and in cells revealed by cryo-electron tomography. Acta Cryst. D 74, 572–584 (2018).

    Article  Google Scholar 

  22. Vitre, B. et al. EB1 regulates microtubule dynamics and tubulin sheet closure in vitro. Nat. Cell Biol. 10, 415–421 (2008).

    Article  Google Scholar 

  23. Doodhi, H. et al. Termination of protofilament elongation by eribulin induces lattice defects that promote microtubule catastrophes. Curr. Biol. 26, 1713–1721 (2016).

    Article  Google Scholar 

  24. Schaap, I. T., de Pablo, P. J. & Schmidt, C. F. Resolving the molecular structure of microtubules under physiological conditions with scanning force microscopy. Eur. Biophys. J. 33, 462–467 (2004).

    Article  Google Scholar 

  25. Weisenberg, R. C. Microtubule formation in vitro in solutions containing low calcium concentrations. Science 177, 1104–1105 (1972).

    Article  ADS  Google Scholar 

  26. Kellogg, E. H. et al. Insights into the distinct mechanisms of action of taxane and non-taxane microtubule stabilizers from cryo-EM structures. J. Mol. Biol. 429, 633–646 (2017).

    Article  Google Scholar 

  27. Yajima, H. et al. Conformational changes in tubulin in GMPCPP and GDP-taxol microtubules observed by cryoelectron microscopy. J. Cell Biol. 198, 315–322 (2012).

    Article  Google Scholar 

  28. Kirkwood, T. B. L. Geometric means and measures of dispersion. Biometrics 35, 908–909 (1979).

    Google Scholar 

  29. Mandelkow, E.-M., Schultheiss, R., Rapp, R., Müller, M. & Mandelkow, E. On the surface lattice of microtubules: helix starts, protofilament number, seam, and handedness. J. Cell Biol. 102, 1067–1073 (1986).

    Article  Google Scholar 

  30. Chrétien, D. & Wade, R. H. New data on the microtubule surface lattice. Biol. Cell 71, 161–174 (1991).

    Article  Google Scholar 

  31. Gardner, M. K. et al. Rapid microtubule self-assembly kinetics. Cell 146, 582–592 (2011).

    Article  Google Scholar 

  32. Wu, Z. et al. Simulations of tubulin sheet polymers as possible structural intermediates in microtubule assembly. PLoS ONE 4, e7291 (2009).

    Article  ADS  Google Scholar 

  33. Hunyadi, V., Chrétien, D. & Jánosi, I. M. Mechanical stress induced mechanism of microtubule catastrophes. J. Mol. Biol. 348, 927–938 (2005).

    Article  Google Scholar 

  34. Janson, M. E. & Dogterom, M. A bending mode analysis for growing microtubules: evidence for a velocity-dependent rigidity. Biophys. J. 87, 2723–2736 (2004).

    Article  ADS  Google Scholar 

  35. Chrétien, D., Fuller, S. D. & Karsenti, E. Structure of growing microtubule ends: two-dimensional sheets close into tubes at variable rates. J. Cell Biol. 129, 1311–1328 (1995).

    Article  Google Scholar 

  36. Coquelle, F. et al. Cryo-electron tomography of microtubules assembled in vitro from purified components. Methods Mol. Biol. 777, 193–208 (2011).

    Article  Google Scholar 

  37. Hyman, A. A., Chrétien, D., Arnal, I. & Wade, R. Structural changes accompanying GTP hydrolysis in microtubules: information from a slowly hydrolyzable anlalogue guanylyl-(α,β)-methylene-diphosphonate. J. Cell Biol. 128, 117–125 (1995).

    Article  Google Scholar 

  38. Shelanski, M. L. Chemistry of the filaments and tubules of brain. J. Histochem. Cytochem. 21, 529–539 (1973).

    Article  Google Scholar 

  39. Malekzadeh-Hemmat, K., Gendry, P. & Launey, J. F. Rat pancreas kinesin: identification and potential binding to microtubules. Cell Mol. Biol. 39, 279–285 (1993).

    Google Scholar 

  40. Hyman, A. et al. Preparation of modified tubulins. Methods Enzymol. 196, 478–485 (1991).

    Article  Google Scholar 

  41. Portran, D., Gaillard, J., Vantard, M. & Théry, M. Quantification of MAP and molecular motor activities on geometrically controlled microtubule networks. Cytoskeleton 70, 12–23 (2013).

    Article  Google Scholar 

  42. Duffy, D. C., McDonald, J. C., Schueller, O. J. & Whitesides, G. M. Rapid prototyping of microfluidic systems in poly(dimethylsiloxane). Anal. Chem. 70, 4974–4984 (1998).

    Article  Google Scholar 

  43. Chrétien, D., Buendia, B., Fuller, S. D. & Karsenti, E. Reconstruction of the centrosome cycle from cryoelectron micrographs. J. Struct. Biol. 120, 117–133 (1997).

    Article  Google Scholar 

  44. Weis, F., Moullintraffort, L., Heichette, C., Chrétien, D. & Garnier, C. The 90-kDa heat shock protein HSP90 protects tubulin against thermal denaturation. J. Biol. Chem. 285, 952–534 (2010).

    Google Scholar 

  45. Mastronarde, D. N. Dual-axis tomography: an approach with alignment methods that preserve resolution. J. Struct. Biol. 120, 343–352 (1997).

    Article  Google Scholar 

  46. Pettersen, E. F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).

    Article  Google Scholar 

  47. Lukkien, J. J., Segers, J. P. L., Hilbers, P. A. J., Gelten, R. J. & Jansen, A. P. J. Efficient Monte Carlo methods for the simulation of catalytic surface reactions. Phys. Rev. E 58, 2598–2610 (1998).

    Article  ADS  Google Scholar 

  48. Sui, H. & Downing, K. H. Structural basis of interprotofilament interaction and lateral deformation of microtubules. Structure 18, 1022–1031 (2010).

    Article  Google Scholar 

  49. Alushin, G. M. et al. High-resolution microtubule structures reveal the structural transitions in αβ-tubulin upon GTP hydrolysis. Cell 157, 1117–1129 (2014).

    Article  Google Scholar 

  50. Groot, De, S. R. & Mazur, P. Non-Equilibrium Thermodynamics (Dover Publications Inc., 1984).

  51. Jmol: an open-source Java viewer for chemical structures in 3D; http://www.jmol.org

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Acknowledgements

This work was supported by the French National Agency for Research (ANR-16-CE11-0017-01 to D.C., ANR-12-BSV5-0004-01 to M.T., ANR-14-CE09-0014-02 to L.B. and ANR-18-CE13-0001 to K.J., M.T. and D.C.), the Human Frontier in Science Program (RGY0088 to M.T.) and the European Research Council (Consolidator Grant 771599 (ICEBERG) to M.T. and Advanced Grant 741773 (AAA) to L.B.).

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Authors and Affiliations

  1. Univ. Grenoble-Alpes, CEA, CNRS, INRA, Interdisciplinary Research Institute of Grenoble, Laboratoire de Physiologie Cellulaire & Végétale, CytoMorpho Lab, Grenoble, France

    Laura Schaedel, Sarah Triclin, Charlotte Aumeier, Jérémie Gaillard, Laurent Blanchoin & Manuel Théry

  2. Univ. Rennes, CNRS, IGDR (Institute of Genetics and Development of Rennes)—UMR 6290, Rennes, France

    Denis Chrétien

  3. CRBM, CNRS, University of Montpellier, Montpellier, France

    Ariane Abrieu

  4. Univ. Paris Diderot, INSERM, CEA, Hôpital Saint Louis, Institut Universitaire d’Hematologie, UMRS 1160, CytoMorpho Lab, Paris, France

    Laurent Blanchoin & Manuel Théry

  5. Univ. Grenoble-Alpes, CNRS, Laboratoire Interdisciplinaire de Physique, Grenoble, France

    Karin John

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  1. Laura Schaedel
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Contributions

L.S. and S.T. performed all dimer exchange and fracture experiments with the help of J.G. and A.A. L.S., L.B. and M.T. designed these experiments. D.C. designed and performed cryo-electron microscopy experiments. K.J. designed and performed numerical simulations. L.S., S.T., C.A., L.B., M.T. and K.J. analysed data. L.S., M.T., D.C. and K.J. wrote the manuscript.

Corresponding authors

Correspondence to Laurent Blanchoin, Manuel Théry or Karin John.

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Supplementary Information, Supplementary Methods, Supplementary Figs. 1–8, Supplementary Tables 1 and 2, and Supplementary References 1–19.

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Schaedel, L., Triclin, S., Chrétien, D. et al. Lattice defects induce microtubule self-renewal. Nat. Phys. 15, 830–838 (2019). https://doi.org/10.1038/s41567-019-0542-4

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