Structural transitions of F-actin upon ATP hydrolysis at near-atomic resolution revealed by cryo-EM

Abstract

The function of actin is coupled to the nucleotide bound to its active site. ATP hydrolysis is activated during polymerization; a delay between hydrolysis and inorganic phosphate (Pi) release results in a gradient of ATP, ADP–Pi and ADP along actin filaments (F-actin). Actin-binding proteins can recognize F-actin’s nucleotide state, using it as a local ‘age’ tag. The underlying mechanism is complex and poorly understood. Here we report six high-resolution cryo-EM structures of F-actin from rabbit skeletal muscle in different nucleotide states. The structures reveal that actin polymerization repositions the proposed catalytic base, His161, closer to the γ-phosphate. Nucleotide hydrolysis and Pi release modulate the conformational ensemble at the periphery of the filament, thus resulting in open and closed states, which can be sensed by coronin-1B. The drug-like toxin jasplakinolide locks F-actin in an open state. Our results demonstrate in detail how ATP hydrolysis links to F-actin’s conformational dynamics and protein interaction.

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Fig. 1: Effect of nucleotide hydrolysis and phosphate release on the intrastrand interface of F-actin.
Fig. 2: Densities of the small molecules bound to F-actin in the different reconstructions.
Fig. 3: Conformational changes at the active site during the G-to-F transition.
Fig. 4: Effect of JASP binding on the structure of F-actin.
Fig. 5: Principal component analysis of the effect of ligand binding on the conformational landscape of F-actin.
Fig. 6: Binding of coronin-1B to F-actin in complex with different ligands, measured by TIRF microscopy.
Fig. 7: Schematic illustration of actin polymerization, nucleotide hydrolysis and the effect of JASP.

References

  1. 1.

    Straub, F. B. & Feuer, G. Adenosine triphosphate, the functional group of actin. Kiserl. Orvostud. 2, 141–151 (1950).

    PubMed  CAS  Google Scholar 

  2. 2.

    Laki, K., Bowen, W. J. & Clark, A. The polymerization of proteins; adenosine triphosphate and the polymerization of actin. J. Gen. Physiol. 33, 437–443 (1950).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  3. 3.

    Combeau, C. & Carlier, M. F. Probing the mechanism of ATP hydrolysis on F-actin using vanadate and the structural analogs of phosphate BeF-3 and A1F-4. J. Biol. Chem. 263, 17429–17436 (1988).

    PubMed  CAS  Google Scholar 

  4. 4.

    Cai, L., Makhov, A. M. & Bear, J. E. F-actin binding is essential for coronin 1B function in vivo. J. Cell Sci. 120, 1779–1790 (2007).

    Article  PubMed  CAS  Google Scholar 

  5. 5.

    Blanchoin, L. & Pollard, T. D. Mechanism of interaction of Acanthamoeba actophorin (ADF/Cofilin) with actin filaments. J. Biol. Chem. 274, 15538–15546 (1999).

    Article  PubMed  CAS  Google Scholar 

  6. 6.

    Suarez, C. et al. Cofilin tunes the nucleotide state of actin filaments and severs at bare and decorated segment boundaries. Curr. Biol. 21, 862–868 (2011).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  7. 7.

    Cai, L., Marshall, T. W., Uetrecht, A. C., Schafer, D. A. & Bear, J. E. Coronin 1B coordinates Arp2/3 complex and cofilin activities at the leading edge. Cell 128, 915–929 (2007).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  8. 8.

    Pollard, T. D. & Borisy, G. G. Cellular motility driven by assembly and disassembly of actin filaments. Cell 112, 453–465 (2003).

    Article  PubMed  CAS  Google Scholar 

  9. 9.

    Blanchoin, L., Pollard, T. D. & Mullins, R. D. Interactions of ADF/cofilin, Arp2/3 complex, capping protein and profilin in remodeling of branched actin filament networks. Curr. Biol. 10, 1273–1282 (2000).

    Article  PubMed  CAS  Google Scholar 

  10. 10.

    Kudryashov, D. S. & Reisler, E. ATP and ADP actin states. Biopolymers 99, 245–256 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  11. 11.

    Oztug Durer, Z. A., Diraviyam, K., Sept, D., Kudryashov, D. S. & Reisler, E. F-actin structure destabilization and DNase I binding loop: fluctuations mutational cross-linking and electron microscopy analysis of loop states and effects on F-actin. J. Mol. Biol. 395, 544–557 (2010).

    Article  PubMed  CAS  Google Scholar 

  12. 12.

    Mannherz, H. G., Brehme, H. & Lamp, U. Depolymerisation of F-actin to G-actin and its repolymerisation in the presence of analogs of adenosine triphosphate. Eur. J. Biochem. 60, 109–116 (1975).

    Article  PubMed  CAS  Google Scholar 

  13. 13.

    Graceffa, P. & Dominguez, R. Crystal structure of monomeric actin in the ATP state: structural basis of nucleotide-dependent actin dynamics. J. Biol. Chem. 278, 34172–34180 (2003).

    Article  PubMed  CAS  Google Scholar 

  14. 14.

    Cooke, R. The role of the bound nucleotide in the polymerization of actin. Biochemistry 14, 3250–3256 (1975).

    Article  PubMed  CAS  Google Scholar 

  15. 15.

    Courtemanche, N. & Pollard, T. D. Interaction of profilin with the barbed end of actin filaments. Biochemistry 52, 6456–6466 (2013). 34, 8960–8972.

    Article  PubMed  CAS  Google Scholar 

  16. 16.

    Fisher, A.J. et al. X-ray structures of the myosin motor domain of Dictyostelium discoideum complexed with MgADP·BeFx and MgADP·AlF4. Biochemistry (1995).

  17. 17.

    Gulick, A. M., Bauer, C. B., Thoden, J. B. & Rayment, I. X-ray structures of the MgADP, MgATPgammaS, and MgAMPPNP complexes of the Dictyostelium discoideum myosin motor domain. Biochemistry 36, 11619–11628 (1997).

    Article  PubMed  CAS  Google Scholar 

  18. 18.

    Rould, M. A., Wan, Q., Joel, P. B., Lowey, S. & Trybus, K. M. Crystal structures of expressed nonpolymerizable monomeric actin in the ADP and ATP states. J. Biol. Chem. 281, 31909–31919 (2006).

    Article  PubMed  CAS  Google Scholar 

  19. 19.

    Vorobiev, S. et al. The structure of nonvertebrate actin: implications for the ATP hydrolytic mechanism. Proc. Natl. Acad. Sci. USA 100, 5760–5765 (2003).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  20. 20.

    McCullagh, M., Saunders, M. G. & Voth, G. A. Unraveling the mystery of ATP hydrolysis in actin filaments. J. Am. Chem. Soc. 136, 13053–13058 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  21. 21.

    Cooke, R. & Murdoch, L. Interaction of actin with analogs of adenosine triphosphate. Biochemistry 12, 3927–3932 (1973).

    Article  PubMed  CAS  Google Scholar 

  22. 22.

    Nolen, B. J. & Pollard, T. D. Insights into the influence of nucleotides on actin family proteins from seven structures of Arp2/3 complex. Mol. Cell 26, 449–457 (2007).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  23. 23.

    Murakami, K. et al. Structural basis for actin assembly, activation of ATP hydrolysis, and delayed phosphate release. Cell 143, 275–287 (2010).

    Article  PubMed  CAS  Google Scholar 

  24. 24.

    Crews, P., Manes, L. V. & Boehler, M. Jasplakinolide, a cyclodepsipeptide from the marine sponge, SP. Tetrahedr. Lett. 27, 2797–2800 (1986).

    CAS  Google Scholar 

  25. 25.

    Bubb, M. R., Spector, I., Beyer, B. B. & Fosen, K. M. Effects of jasplakinolide on the kinetics of actin polymerization: an explanation for certain in vivo observations. J. Biol. Chem. 275, 5163–5170 (2000).

    Article  PubMed  CAS  Google Scholar 

  26. 26.

    Vig, A. et al. The effect of toxins on inorganic phosphate release during actin polymerization. Eur. Biophys. J. 40, 619–626 (2011).

    Article  PubMed  CAS  Google Scholar 

  27. 27.

    Tannert, R. et al. Synthesis and structure-activity correlation of natural-product inspired cyclodepsipeptides stabilizing F-actin. J. Am. Chem. Soc. 132, 3063–3077 (2010).

    Article  PubMed  CAS  Google Scholar 

  28. 28.

    Milroy, L.-G. et al. Selective chemical imaging of static actin in live cells. J. Am. Chem. Soc. 134, 8480–8486 (2012).

    Article  PubMed  CAS  Google Scholar 

  29. 29.

    Lukinavičius, G. et al. Fluorogenic probes for live-cell imaging of the cytoskeleton. Nat. Methods 11, 731–733 (2014).

    Article  PubMed  CAS  Google Scholar 

  30. 30.

    Pospich, S. et al. Near-atomic structure of jasplakinolide-stabilized malaria parasite F-actin reveals the structural basis of filament instability. Proc. Natl. Acad. Sci. USA https://doi.org/10.1073/pnas.1707506114 (2017).

  31. 31.

    Galkin, V. E., Orlova, A., Vos, M. R., Schröder, G. F. & Egelman, E. H. Near-atomic resolution for one state of F-actin. Structure 23, 173–182 (2015).

    Article  PubMed  CAS  Google Scholar 

  32. 32.

    von der Ecken, J. et al. Structure of the F-actin–tropomyosin complex. Nature 519, 114–117 (2015).

    Article  PubMed  CAS  Google Scholar 

  33. 33.

    von der Ecken, J., Heissler, S. M., Pathan-Chhatbar, S., Manstein, D. J. & Raunser, S. Cryo-EM structure of a human cytoplasmic actomyosin complex at near-atomic resolution. Nature 534, 724–728 (2016).

    Article  PubMed  CAS  Google Scholar 

  34. 34.

    Otterbein, L. R., Graceffa, P. & Dominguez, R. The crystal structure of uncomplexed actin in the ADP state. Science 293, 708–711 (2001).

    Article  PubMed  CAS  Google Scholar 

  35. 35.

    Zheng, X., Diraviyam, K. & Sept, D. Nucleotide effects on the structure and dynamics of actin. Biophys. J. 93, 1277–1283 (2007).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  36. 36.

    Isambert, H. et al. Flexibility of actin filaments derived from thermal fluctuations: effect of bound nucleotide, phalloidin, and muscle regulatory proteins. J. Biol. Chem. 270, 11437–11444 (1995).

    Article  PubMed  CAS  Google Scholar 

  37. 37.

    Kardos, R. et al. The effect of jasplakinolide on the thermodynamic properties of ADP.BeF(x) bound actin filaments. Thermochim. Acta 463, 77–80 (2007).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  38. 38.

    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  PubMed  PubMed Central  CAS  Google Scholar 

  39. 39.

    Bharat, T. A. M., Murshudov, G. N., Sachse, C. & Löwe, J. Structures of actin-like ParM filaments show architecture of plasmid-segregating spindles. Nature 523, 106–110 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  40. 40.

    Ge, P., Durer, Z. A. O., Kudryashov, D., Zhou, Z. H. & Reisler, E. Cryo-EM reveals different coronin binding modes for ADP– and ADP–BeFx actin filaments. Nat. Struct. Mol. Biol. 21, 1075–1081 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  41. 41.

    Oda, T., Iwasa, M., Aihara, T., Maéda, Y. & Narita, A. The nature of the globular- to fibrous-actin transition. Nature 457, 441–445 (2009).

    Article  PubMed  CAS  Google Scholar 

  42. 42.

    Barad, B. A. et al. EMRinger: side chain-directed model and map validation for 3D cryo-electron microscopy. Nat. Methods 12, 943–946 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  43. 43.

    Allegretti, M., Mills, D. J., McMullan, G., Kühlbrandt, W. & Vonck, J. Atomic model of the F420-reducing [NiFe] hydrogenase by electron cryo-microscopy using a direct electron detector. eLife 3, e01963 (2014).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  44. 44.

    Wriggers, W. & Schulten, K. Investigating a back door mechanism of actin phosphate release by steered molecular dynamics. Proteins 35, 262–273 (1999).

    Article  PubMed  CAS  Google Scholar 

  45. 45.

    Bubb, M. R., Senderowicz, A. M., Sausville, E. A., Duncan, K. L. & Korn, E. D. Jasplakinolide, a cytotoxic natural product, induces actin polymerization and competitively inhibits the binding of phalloidin to F-actin. J. Biol. Chem. 269, 14869–14871 (1994).

    PubMed  CAS  Google Scholar 

  46. 46.

    Papp, G. et al. Conformational changes in actin filaments induced by formin binding to the barbed end. Biophys. J. 91, 2564–2572 (2006).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  47. 47.

    Strzelecka-Gołaszewska, H., Mossakowska, M., Woźniak, A., Moraczewska, J. & Nakayama, H. Long-range conformational effects of proteolytic removal of the last three residues of actin. Biochem. J. 307, 527–534 (1995).

    Article  PubMed  PubMed Central  Google Scholar 

  48. 48.

    Zimmermann, D., Santos, A., Kovar, D. R. & Rock, R. S. Actin age orchestrates myosin-5 and myosin-6 run lengths. Curr. Biol. 25, 2057–2062 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  49. 49.

    Mentes, A. et al. High-resolution cryo-EM structures of actin-bound myosin states reveal the mechanism of myosin force sensing. Proc. Natl. Acad. Sci. USA 115, 1292–1297 (2018).

    Article  PubMed  CAS  Google Scholar 

  50. 50.

    Galkin, V. E. et al. Remodeling of actin filaments by ADF/cofilin proteins. Proc. Natl. Acad. Sci. USA 108, 20568–20572 (2011).

    Article  PubMed  PubMed Central  Google Scholar 

  51. 51.

    Muhlrad, A., Pavlov, D., Peyser, Y. M. & Reisler, E. Inorganic phosphate regulates the binding of cofilin to actin filaments. FEBS J. 273, 1488–1496 (2006).

    Article  PubMed  CAS  Google Scholar 

  52. 52.

    Moriyama, K. & Yahara, I. The actin-severing activity of cofilin is exerted by the interplay of three distinct sites on cofilin and essential for cell viability. Biochem. J. 365, 147–155 (2002).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  53. 53.

    Kabsch, W., Mannherz, H. G., Suck, D., Pai, E. F. & Holmes, K. C. Atomic structure of the actin:DNase I complex. Nature 347, 37–44 (1990).

    Article  PubMed  CAS  Google Scholar 

  54. 54.

    Aitken, C. E., Marshall, R. A. & Puglisi, J. D. An oxygen scavenging system for improvement of dye stability in single-molecule fluorescence experiments. Biophys. J. 94, 1826–1835 (2008).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  55. 55.

    Rasnik, I., McKinney, S. A. & Ha, T. Nonblinking and long-lasting single-molecule fluorescence imaging: it’s ProQuest. Nat. Methods 3, 891–893 (2006).

    Article  PubMed  CAS  Google Scholar 

  56. 56.

    Bieling, P., Telley, I. A., Hentrich, C., Piehler, J. & Surrey, T. Fluorescence microscopy assays on chemically functionalized surfaces for quantitative imaging of microtubule, motor, and +TIP dynamics. Methods Cell Biol. 95, 555–580 (2010).

    Article  PubMed  CAS  Google Scholar 

  57. 57.

    Pardee, J. D. & Spudich, J. A. Purification of muscle actin. Methods Enzymol. 85, 164–181 (1982).

    Article  PubMed  CAS  Google Scholar 

  58. 58.

    Hansen, S. D., Zuchero, J. B. & Mullins, R. D. Cytoplasmic actin: purification and single molecule assembly assays. Methods Mol. Biol. 1046, 145–170 (2013).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  59. 59.

    Margossian, S. S. & Lowey, S. Preparation of myosin and its subfragments from rabbit skeletal muscle. Methods Enzymol. 85, 55–71 (1982).

    Article  PubMed  CAS  Google Scholar 

  60. 60.

    Pollard, T. D. Myosin purification and characterization. Methods Cell Biol. 24, 333–371 (1982).

    Article  PubMed  CAS  Google Scholar 

  61. 61.

    Zheng, S. Q. et al. MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy. Nat. Methods 14, 331–332 (2017).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  62. 62.

    Grant, T. & Grigorieff, N. Measuring the optimal exposure for single particle cryo-EM using a 2.6 Å reconstruction of rotavirus VP6. eLife 4, e06980 (2015).

    Article  PubMed  PubMed Central  Google Scholar 

  63. 63.

    Zhang, K. Gctf: real-time CTF determination and correction. J. Struct. Biol. 193, 1–12 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  64. 64.

    Rohou, A. & Grigorieff, N. CTFFIND4: fast and accurate defocus estimation from electron micrographs. J. Struct. Biol. 192, 216–221 (2015).

    Article  PubMed  Google Scholar 

  65. 65.

    Behrmann, E. et al. Real-space processing of helical filaments in SPARX. J. Struct. Biol. 177, 302–313 (2012).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  66. 66.

    Scheres, S. H. W. RELION: implementation of a Bayesian approach to cryo-EM structure determination. J. Struct. Biol. 180, 519–530 (2012).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  67. 67.

    Tang, G. et al. EMAN2: an extensible image processing suite for electron microscopy. J. Struct. Biol. 157, 38–46 (2007).

    Article  PubMed  CAS  Google Scholar 

  68. 68.

    Sachse, C. et al. High-resolution electron microscopy of helical specimens: a fresh look at tobacco mosaic virus. J. Mol. Biol. 371, 812–835 (2007).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  69. 69.

    Moriya, T. et al. High-resolution single particle analysis from electron cryo-microscopy images using SPHIRE. J. Vis. Exp. e55448 (2017).

  70. 70.

    Song, Y. et al. High-resolution comparative modeling with RosettaCM. Structure 21, 1735–1742 (2013).

    Article  PubMed  CAS  Google Scholar 

  71. 71.

    Trabuco, L. G., Villa, E., Mitra, K., Frank, J. & Schulten, K. Flexible fitting of atomic structures into electron microscopy maps using molecular dynamics. Structure 16, 673–683 (2008).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  72. 72.

    Huang, J. et al. CHARMM36m: an improved force field for folded and intrinsically disordered proteins. Nat. Methods 14, 71–73 (2017).

    Article  PubMed  CAS  Google Scholar 

  73. 73.

    Phillips, J. C. et al. Scalable molecular dynamics with NAMD. J. Comput. Chem. 26, 1781–1802 (2005).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  74. 74.

    Humphrey, W., Dalke, A. & Schulten, K. VMD: visual molecular dynamics. J. Mol. Graph. 14, 33–38 (1996).

    Article  PubMed  CAS  Google Scholar 

  75. 75.

    Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D Biol. Crystallogr. 66, 486–501 (2010).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  76. 76.

    DiMaio, F. et al. Atomic-accuracy models from 4.5-Å cryo-electron microscopy data with density-guided iterative local refinement. Nat. Methods 12, 361–365 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  77. 77.

    Fleishman, S. J. et al. RosettaScripts: a scripting language interface to the Rosetta macromolecular modeling suite. PLoS One 6, e20161 (2011).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  78. 78.

    Chen, V. B. et al. MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr. D Biol. Crystallogr. 66, 12–21 (2010).

    Article  PubMed  CAS  Google Scholar 

  79. 80.

    Kortemme, T., Kim, D.E. & Baker, D. Computational alanine scanning of protein-protein interfaces. Sci. STKE 2004, pl2 (2004).

  80. 81.

    Abraham, M. J. et al. GROMACS: high performance molecular simulations through multi-level parallelism from laptops to supercomputers. SoftwareX 1–2, 19–25 (2015).

    Article  Google Scholar 

  81. 82.

    Appleton, B. A., Wu, P. & Wiesmann, C. The crystal structure of murine coronin-1: a regulator of actin cytoskeletal dynamics in lymphocytes. Structure 14, 87–96 (2006).

    Article  PubMed  CAS  Google Scholar 

  82. 83.

    Eswar, N., Eramian, D., Webb, B., Shen, M.-Y. & Sali, A. Protein structure modeling with MODELLER. Methods Mol. Biol. 426, 145–159 (2008).

    Article  PubMed  CAS  Google Scholar 

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Acknowledgements

We thank O. Hofnagel and D. Prumbaum for assistance with data collection. We thank W. Linke and A. Unger (Ruhr-Universität Bochum, Germany) for providing us with muscle acetone powder. This work was supported by the Max Planck Society (to S.R.), the state of Thuringia (to H.-D.A., grant 43-5572-321-12040-12) and the European Council under the European Union’s Seventh Framework Programme (FP7/ 2007–2013) (grant 615984) (to S.R.). S.P. is supported as a fellow of Studienstiftung des deutschen Volkes.

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Contributions

S.R. designed the project. F.M. and S.P. prepared and processed cryo-EM specimens, and built the atomic models. T.W. designed and implemented the filament autopicking tool. J.F. and P.B. designed and analyzed the TIRF microscopy experiments. J.F. produced recombinant coronin-1B and performed the TIRF microscopy experiments. F.K. and H.-D.A. synthesized JASP. F.M., S.P. and J.F. prepared figures and videos. F.M. and S.R. wrote the manuscript. All authors reviewed the results and commented on the manuscript.

Corresponding author

Correspondence to Stefan Raunser.

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Integrated Supplementary Information

Supplementary Figure 1 Schematic representation of the modified single-particle analysis method used to obtain the F-actin models.

For each step, possible filament segment alignments are represented along with the development of the Fourier shell correlation curves and the maps with their corresponding distribution of angular assignments. For details see Methods.

Supplementary Figure 2 Overview of the cryo-EM data and resolution for all six datasets.

(a,b) Representative micrographs at ~ -1.5 μm defocus (a) and their power spectrum (b). (c) The local resolution of all reconstructions decreases gradually towards the ends of the filament, reflecting the intrinsic flexibility of F-actin. In addition, SD1 has systematically lower local resolution than the rest of the monomer, suggesting this area is flexible in the filament. The same color scale was used for all volumes. (d) Fourier shell correlation (FSC) curves for the masked (black) and unmasked (green) maps. The FSC plots for the unmasked maps clearly show that there is no over-refinement in the reconstructions. FSCs were calculated for a 120-Å-long central section of the filament. The red curves show the correlation for phase-randomized maps, proving that the masking has no effect on the estimated resolution.

Supplementary Figure 3 Density at the nucleotide-binding site.

Stereo pair images for the nucleotide-binding site of F-actin in complex with AppNHp (a), ADP–BeFx (b), ADP–Pi (c), ADP–Pi JASP (d), ADP JASP (e), and ADP (f). The images show the density for the metal-nucleotide complex (yellow mesh) and protein (transparent gray). In addition, we included several key amino acids at the nucleotide-binding pocket. For further orientation and residue labeling please see Fig. 2.

Supplementary Figure 4 G-to F transition of AppNHp-actin.

The superposition of the models of AppNHp-actin in the G- (blue, bound Ca2+, PDB 1NWK (Graceffa, P., and Dominguez, R., J Biol Chem. 278, 34172–34180, 2003)) and F-state (red, bound Mg2+) at the center illustrates the conformational change upon polymerization. Insets show the nucleotide-binding pockets of G-actin (left) and F-actin (right) including possible hydrogen bonds between the nucleotide and the protein, highlighted by green dashed lines. The interactions of the γ-phosphate with the protein backbone found in G-actin are probably preserved in the F-state. Both models are shown as side (upper panel) and top (lower panel) views (see also Supplementary Video 3).

Supplementary Figure 5 Evaluation of the nucleotide composition of the AppNHp filaments by reverse-phase ion-pair chromatography.

Before grid preparation, filaments were collected and resuspended in F-buffer without nucleotides. Samples were heat-denatured and loaded directly onto the column. Shaded regions represent peaks coming from AppNHp and AppNH2 (blue), ADP (turquoise), or the buffer (gray). The fractions of total nucleotides in the sample are indicated for each species, demonstrating that the AppNHp filaments contained a mixture of AppNHp and its hydrolysis products. Buffer control represents a run with F-buffer, necessary to identify the signal of the buffer components.

Supplementary Figure 6 Comparison between bare F-actin–ADP and its complex with tropomyosin.

(a) Superposition of the atomic models of F-actin–ADP (turquoise) and F-actin–ADP–tropomyosin (beige and green, PDB 5JLF, EMDB 8162 (von der Ecken, J., et al., Nature. 534, 724–728, 2016; von der Ecken, J., et al., Nature. 519, 114–117, 2015). The C terminal tails of all F-actin subunits are highlighted in red. (b, c) Close-up views of the C termini and corresponding densities in the absence (b) and presence (c) of tropomyosin, illustrating that tropomyosin binding results in the disordering of the C terminal tail.

Supplementary Figure 7 In silico alanine scanning for F-actin in the open and closed D loop conformations.

The interface was defined based on the central actin subunit of each model filament. (a) Predicted change of free energy upon alanine mutations in the closed (represented by F-actin–ADP, top, turquoise) and open (represented by F-actin–ADP–BeFx, bottom, orange) conformations of the intrastrand interface. The dotted lines indicate the threshold over which alanine mutations are considered destabilizing. (b) Predicted effect of the alanine mutations mapped in the filament structures of F-actin–ADP (closed state, top) and F-actin–ADP–BeFx (open state, bottom). In the open state, the effect is generally bigger than in the closed state, explaining the lower stability of F-actin–ADP. For guidance, regions forming the filament interfaces are highlighted and labeled, these include the D loop (a.a. 38 – 52), proline rich loop (P loop, a.a. 108 – 112), W loop (a.a. 165 – 172), plug (H plug, a.a. 263 – 273), and the C terminus. Subunits are shown as surfaces calculated from backbone and Cβ-atoms, except the central one, which is shown in ribbons. The color scale represents the predicted change of free energy from destabilizing (red) to stabilizing (blue).

Supplementary Figure 8 Proposed effect of the nucleotide state of actin on the binding of different actin-binding proteins.

(a-c) Molecular dynamics flexible fitting models of the complex between yeast coronin and F-actin. Ribbon representation of F-actin in complex with ADP–BeFx (red, a) or ADP states (green, b) complexed with coronin (blue). The insets show possible contacts between the D loop and coronin residues, illustrating that coronin could directly sense the changes occurring in this region. (c) To exclude a possible direct interaction of JASP with coronin, we superimposed the density found in the JASP complexes (yellow) with the ADP–BeFx filament. The results highlight the lack of overlap between the coronin and JASP binding sites. The molecular dynamics flexible fitting was performed using previously published electron density maps (EMDB accession codes 6101 and 6100, respectively (Ge, P., et al., Nat Struct Mol Biol. 21, 1075–1081, 2014.)). (d,e) Superposition between the F-actin–cofilin (gray/purple) complex (PDB 3J0S (Galkin, V.E., et al., Proc Nat Acad Sci. 108, 20568–20572, 2011)) and our ADP–BeFx (red, d) and ADP (green, e) models. The first 5 residues of cofilin, shown to be sensitive to the nucleotide state of F-actin (Moriyama, K., and Yahara, I., Biochem J. 365, 147–155, 2002) are highlighted in yellow. The figure illustrates how the N terminus of cofilin binds to the same pocket as the C terminus of actin in the F-actin–ADP–BeFx structure (d). In addition, the position of S3 is highlighted, which is known to be phosphorylated to regulate the function of cofilin.

Supplementary information

Supplementary Text and Figures

Supplementary Figures 1–8 and Supplementary Note

Reporting Summary

Supplementary Video 1

Agreement between models and maps. (a–f) Model-map agreement of the (a) F-actin–AppNHp, (b) F-actin–ADP–BeFx, (c) F-actin–ADP–Pi, (d) F-actin–ADP–Pi JASP, (e) F-actin–ADP JASP, and (f) F-actin–ADP reconstructions. For each map the same representative α-helical and β-sheet regions as well as the nucleotide-binding pocket are shown. Ribbons are colored according to the nucleotide state represented in the model: ATP-like: AppNHp/ADP–BeFx (blue), ADP–Pi-like: ADP–Pi/ADP–Pi JASP (red) and ADP-like: ADP/ADP JASP (green). The central subunits are colored in a lighter shade to highlight the dimensions of a single subunit. Maps are depicted at comparable thresholds.

Supplementary Video 2

Density at the ligand binding sites. (a–f) The movie shows the density for the ligand and key amino acids at the nucleotide-binding sites of F-actin–AppNHp (a), F-actin–ADP–BeFx (b), F-actin–ADP–Pi (c), F-actin–ADP–Pi JASP (d), F-actin–ADP JASP (e), and F-actin–ADP (f). For F-actin–ADP–BeFx, the density corresponding to BeFx is highlighted in magenta. In addition, the density for JASP and key binding residues are also shown for F-actin–ADP–Pi JASP (d) and F-actin–ADP JASP (e).

Supplementary Video 3

G- to F-actin transition at the active center of AppNHp-bound actin. (a) Whole monomer conformational change. Residues Q137, D154, and H161 as well as the AppNHp and Mg2+ are shown in balls and sticks. Only residues present in the G- (PDB-ID: 1NWK) and F-actin model can be included in the morph, therefore residues 40 – 51 (D loop) and 372-375 (C terminus) are not shown. (b) Close-up view of the conformational change. The density corresponds to the AppNHp map. The morph demonstrates that the new conformation of His161 is supported by the data. (c) Overall fit of the atomic model to the sharpened cryo-EM density.

Supplementary Video 4

Intrastrand interactions at the D loop/C terminus interface. (af) Visualization of side chains and their corresponding densities at the D loop/C terminus interface of the (a) F-actin–AppNHp, (b) F-actin–ADP–BeFx, (c) F-actin–ADP–Pi, (d) F-actin–ADP–Pi JASP, (e) F-actin–ADP JASP, and (f) F-actin–ADP reconstructions. The video highlights the different intrastrand interactions in association to the nucleotide states. All maps were filtered to their local resolution. The interface is located near the center of the reconstruction, with the central subunit shown in a lighter shade.

Supplementary Video 5

Mixed conformations at the D loop/C terminus interface of the F-actin–AppNHp, ADP–BeFx and ADP–Pi map. (a) The sharpened map of F-actin–AppNHp (gray) is superimposed with the corresponding atomic model (shades of blue). (b) Close-up view of the D loop and C terminus interface of F-actin–AppNHp map filtered to 4.1 Å. The D loop is not included in the model, as it is disordered when the map is filtered to the nominal resolution. The density of the C terminus splits, resembling two distinct conformations. Superimposition of the F-actin–AppNHp map (gray) with the model and map of F-actin–ADP (close D loop state, shades of green, c) and F-actin–ADP–Pi JASP (open D loop state, shades of red, d) illustrating a mixture of conformations at the D loop/C terminus interface of F-actin–AppNHp. (e–h) and (i–l) show the same views as in (a–d) but for F-actin–ADP–BeFx (shades of blue) and F-actin–ADP–Pi (shades of red), respectively. Maps shown in (b–d, f–h and j–l) were filtered to 4.1 Å, corresponding to the nominal resolution of the F-actin–ADP map, and are shown at lower thresholds to visualize conformations with low occupancy.

Supplementary Video 6

Principal component analysis of the different actin structures. The movie shows the variation of an F-actin monomer projected on either the first or the second principal component. The dotted line in the plot (left) illustrates the position of the structure (right) along the corresponding component. For clarity, two views are provided for each projection. We used only Cα atoms present in all six structures for the analysis, so part of the D loop (a.a. 45-49) and the C terminus (a.a. 372-375) are not included in the movie. The actin monomer is colored according to its subdomain structure; SD1 is cyan, SD2 green, SD3 orange, and SD4 red.

Supplementary Video 7

Conformational change at the D-loop/C-terminus interface. (a) The sharpened map of F-actin–ADP–Pi JASP (gray) is superimposed with the corresponding atomic model (shades of red) with JASP highlighted in yellow. (b) Close-up view of the D loop and C terminal interface of F-actin–ADP–Pi JASP. (c) Morph (gray) of atomic models illustrating the conformational change of the D loop and C terminus from the open to the closed state. (d) Close-up view of the F-actin–ADP map (gray) and the corresponding model (shades of green). (e) Overview of the complete sharpened F-actin–ADP map and model. Maps shown in (b,d) were filtered to their local resolution.

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Merino, F., Pospich, S., Funk, J. et al. Structural transitions of F-actin upon ATP hydrolysis at near-atomic resolution revealed by cryo-EM. Nat Struct Mol Biol 25, 528–537 (2018). https://doi.org/10.1038/s41594-018-0074-0

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