Structure of the inner kinetochore CCAN complex assembled onto a centromeric nucleosome

Article metrics

Abstract

In eukaryotes, accurate chromosome segregation in mitosis and meiosis maintains genome stability and prevents aneuploidy. Kinetochores are large protein complexes that, by assembling onto specialized Cenp-A nucleosomes1,2, function to connect centromeric chromatin to microtubules of the mitotic spindle3,4. Whereas the centromeres of vertebrate chromosomes comprise millions of DNA base pairs and attach to multiple microtubules, the simple point centromeres of budding yeast are connected to individual microtubules5,6. All 16 budding yeast chromosomes assemble complete kinetochores using a single Cenp-A nucleosome (Cenp-ANuc), each of which is perfectly centred on its cognate centromere7,8,9. The inner and outer kinetochore modules are responsible for interacting with centromeric chromatin and microtubules, respectively. Here we describe the cryo-electron microscopy structure of the Saccharomyces cerevisiae inner kinetochore module, the constitutive centromere associated network (CCAN) complex, assembled onto a Cenp-A nucleosome (CCAN–Cenp-ANuc). The structure explains the interdependency of the constituent subcomplexes of CCAN and shows how the Y-shaped opening of CCAN accommodates Cenp-ANuc to enable specific CCAN subunits to contact the nucleosomal DNA and histone subunits. Interactions with the unwrapped DNA duplex at the two termini of Cenp-ANuc are mediated predominantly by a DNA-binding groove in the Cenp-L–Cenp-N subcomplex. Disruption of these interactions impairs assembly of CCAN onto Cenp-ANuc. Our data indicate a mechanism of Cenp-A nucleosome recognition by CCAN and how CCAN acts as a platform for assembly of the outer kinetochore to link centromeres to the mitotic spindle for chromosome segregation.

Access options

Rent or Buy article

Get time limited or full article access on ReadCube.

from$8.99

All prices are NET prices.

Fig. 1: Structure of the S. cerevisiae CCAN complex.
Fig. 2: Structure of the S. cerevisiae CCAN–Cenp-ANuc complex.
Fig. 3: Cenp-LN interacts with the unwrapped DNA duplex of Cenp-ANuc.
Fig. 4: The Cenp-N DNA-binding groove is required for stable CCAN–Cenp-ANuc interactions.

Data availability

Electron microscopy maps have been deposited with the Electron Microscopy Data Bank with accession codes EMD-4580 (CCAN), EMD-4579 (CCAN-Cenp-ANuc), EMD-4581 (mask1) and EMD-4971 (mask2). Protein coordinates have been deposited with the PDB with accession codes 6QLE (CCAN), 6QLD (CCAN-Cenp-ANuc) and 6QLF (mask1). The XL-MS raw files, the associated output and databases have been deposited through the ProteomeXchange Consortium via the PRIDE partner repository with the dataset identifier PXD013769. Other data are available upon reasonable request.

References

  1. 1.

    Earnshaw, W. C. & Rothfield, N. Identification of a family of human centromere proteins using autoimmune sera from patients with scleroderma. Chromosoma 91, 313–321 (1985).

  2. 2.

    Meluh, P. B., Yang, P., Glowczewski, L., Koshland, D. & Smith, M. M. Cse4p is a component of the core centromere of Saccharomyces cerevisiae. Cell 94, 607–613 (1998).

  3. 3.

    Cheeseman, I. M. The kinetochore. Cold Spring Harb. Perspect. Biol. 6, a015826 (2014).

  4. 4.

    Musacchio, A. & Desai, A. A molecular view of kinetochore assembly and function. Biology 6, 5 (2017).

  5. 5.

    Clarke, L. & Carbon, J. Isolation of a yeast centromere and construction of functional small circular chromosomes. Nature 287, 504–509 (1980).

  6. 6.

    Winey, M. et al. Three-dimensional ultrastructural analysis of the Saccharomyces cerevisiae mitotic spindle. J. Cell Biol. 129, 1601–1615 (1995).

  7. 7.

    Furuyama, S. & Biggins, S. Centromere identity is specified by a single centromeric nucleosome in budding yeast. Proc. Natl Acad. Sci. USA 104, 14706–14711 (2007).

  8. 8.

    Camahort, R. et al. Cse4 is part of an octameric nucleosome in budding yeast. Mol. Cell 35, 794–805 (2009).

  9. 9.

    Lang, J., Barber, A. & Biggins, S. An assay for de novo kinetochore assembly reveals a key role for the CENP-T pathway in budding yeast. eLife 7, e37819 (2018).

  10. 10.

    Hinshaw, S. M. & Harrison, S. C. The structure of the Ctf19c/CCAN from budding yeast. eLife 8, e44239 (2019).

  11. 11.

    McKinley, K. L. et al. The CENP-L–N complex forms a critical node in an integrated meshwork of interactions at the centromere-kinetochore interface. Mol. Cell 60, 886–898 (2015).

  12. 12.

    Pesenti, M. E. et al. Reconstitution of a 26-subunit human kinetochore reveals cooperative microtubule binding by CENP-OPQUR and NDC80. Molecular Cell 71, 923–939 (2018).

  13. 13.

    Schmitzberger, F. & Harrison, S. C. RWD domain: a recurring module in kinetochore architecture shown by a Ctf19–Mcm21 complex structure. EMBO Rep. 13, 216–222 (2012).

  14. 14.

    Dimitrova, Y. N., Jenni, S., Valverde, R., Khin, Y. & Harrison, S. C. Structure of the MIND complex defines a regulatory focus for yeast kinetochore assembly. Cell 167, 1014–1027 (2016).

  15. 15.

    Petrovic, A. et al. Structure of the MIS12 complex and molecular basis of its interaction with CENP-C at human kinetochores. Cell 167, 1028–1040 (2016).

  16. 16.

    Hu, L. et al. Structural analysis of fungal CENP-H/I/K homologs reveals a conserved assembly mechanism underlying proper chromosome alignment. Nucleic Acids Res. 47, 468–479 (2019).

  17. 17.

    Pekgöz Altunkaya, G. et al. CCAN assembly configures composite binding interfaces to promote cross-linking of Ndc80 complexes at the kinetochore. Curr. Biol. 26, 2370–2378 (2016).

  18. 18.

    Guse, A., Carroll, C. W., Moree, B., Fuller, C. J. & Straight, A. F. In vitro centromere and kinetochore assembly on defined chromatin templates. Nature 477, 354–358 (2011).

  19. 19.

    Carroll, C. W., Silva, M. C., Godek, K. M., Jansen, L. E. & Straight, A. F. Centromere assembly requires the direct recognition of CENP-A nucleosomes by CENP-N. Nat. Cell Biol. 11, 896–902 (2009).

  20. 20.

    Carroll, C. W., Milks, K. J. & Straight, A. F. Dual recognition of CENP-A nucleosomes is required for centromere assembly. J. Cell Biol. 189, 1143–1155 (2010).

  21. 21.

    Kingston, I. J., Yung, J. S. & Singleton, M. R. Biophysical characterization of the centromere-specific nucleosome from budding yeast. J. Biol. Chem. 286, 4021–4026 (2011).

  22. 22.

    Tachiwana, H. et al. Crystal structure of the human centromeric nucleosome containing CENP-A. Nature 476, 232–235 (2011).

  23. 23.

    Roulland, Y. et al. The flexible ends of CENP-A nucleosome are required for mitotic fidelity. Mol. Cell 63, 674–685 (2016).

  24. 24.

    White, C. L., Suto, R. K. & Luger, K. Structure of the yeast nucleosome core particle reveals fundamental changes in internucleosome interactions. EMBO J. 20, 5207–5218 (2001).

  25. 25.

    Nishino, T. et al. CENP-T–W–S–X forms a unique centromeric chromatin structure with a histone-like fold. Cell 148, 487–501 (2012).

  26. 26.

    Hornung, P. et al. A cooperative mechanism drives budding yeast kinetochore assembly downstream of CENP-A. J. Cell Biol. 206, 509–524 (2014).

  27. 27.

    Anedchenko, E. A. et al. The kinetochore module Okp1CENP-Q/Ame1CENP-U is a reader for N-terminal modifications on the centromeric histone Cse4CENP-A. EMBO J. 38, e98991 (2019).

  28. 28.

    Chittori, S. et al. Structural mechanisms of centromeric nucleosome recognition by the kinetochore protein CENP-N. Science 359, 339–343 (2018).

  29. 29.

    Pentakota, S. et al. Decoding the centromeric nucleosome through CENP-N. eLife 6, e33442 (2017).

  30. 30.

    Kato, H. et al. A conserved mechanism for centromeric nucleosome recognition by centromere protein CENP-C. Science 340, 1110–1113 (2013).

  31. 31.

    Kouprina, N. et al. Identification and cloning of the CHL4 gene controlling chromosome segregation in yeast. Genetics 135, 327–341 (1993).

  32. 32.

    Weir, J. R. et al. Insights from biochemical reconstitution into the architecture of human kinetochores. Nature 537, 249–253 (2016).

  33. 33.

    Falk, S. J. et al. Chromosomes. CENP-C reshapes and stabilizes CENP-A nucleosomes at the centromere. Science 348, 699–703 (2015).

  34. 34.

    Zhang, Z., Yang, J. & Barford, D. Recombinant expression and reconstitution of multiprotein complexes by the USER cloning method in the insect cell-baculovirus expression system. Methods 95, 13–25 (2016).

  35. 35.

    Schuck, P. On the analysis of protein self-association by sedimentation velocity analytical ultracentrifugation. Anal. Biochem. 320, 104–124 (2003).

  36. 36.

    Brautigam, C. A. Calculations and publication-quality illustrations for analytical ultracentrifugation data. Methods Enzymol. 562, 109–133 (2015).

  37. 37.

    Samel, A., Cuomo, A., Bonaldi, T. & Ehrenhofer-Murray, A. E. Methylation of CenH3 arginine 37 regulates kinetochore integrity and chromosome segregation. Proc. Natl Acad. Sci. USA 109, 9029–9034 (2012).

  38. 38.

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

  39. 39.

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

  40. 40.

    Fernandez-Leiro, R. & Scheres, S. H. W. A pipeline approach to single-particle processing in RELION. Acta Crystallogr. D 73, 496–502 (2017).

  41. 41.

    Nakane, T., Kimanius, D., Lindahl, E. & Scheres, S. H. Characterisation of molecular motions in cryo-EM single-particle data by multi-body refinement in RELION. eLife 7, e36861 (2018).

  42. 42.

    Elmlund, H., Elmlund, D. & Bengio, S. PRIME: probabilistic initial 3D model generation for single-particle cryo-electron microscopy. Structure 21, 1299–1306 (2013).

  43. 43.

    Chen, S. et al. High-resolution noise substitution to measure overfitting and validate resolution in 3D structure determination by single particle electron cryomicroscopy. Ultramicroscopy 135, 24–35 (2013).

  44. 44.

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

  45. 45.

    Yang, Z. et al. UCSF Chimera, MODELLER, and IMP: an integrated modeling system. J. Struct. Biol. 179, 269–278 (2012).

  46. 46.

    Schmitzberger, F. et al. Molecular basis for inner kinetochore configuration through RWD domain–peptide interactions. EMBO J. 36, 3458–3482 (2017).

  47. 47.

    Hinshaw, S. M. & Harrison, S. C. An Iml3–Chl4 heterodimer links the core centromere to factors required for accurate chromosome segregation. Cell Rep. 5, 29–36 (2013).

  48. 48.

    Kelley, L. A., Mezulis, S., Yates, C. M., Wass, M. N. & Sternberg, M. J. The Phyre2 web portal for protein modeling, prediction and analysis. Nat. Protoc. 10, 845–858 (2015).

  49. 49.

    Buchan, D. W., Minneci, F., Nugent, T. C., Bryson, K. & Jones, D. T. Scalable web services for the PSIPRED protein analysis workbench. Nucleic Acids Res. 41, W349–W357 (2013).

  50. 50.

    Luger, K., Mäder, A. W., Richmond, R. K., Sargent, D. F. & Richmond, T. J. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389, 251–260 (1997).

  51. 51.

    Vasudevan, D., Chua, E. Y. D. & Davey, C. A. Crystal structures of nucleosome core particles containing the ‘601’ strong positioning sequence. J. Mol. Biol. 403, 1–10 (2010).

  52. 52.

    Adams, P. D. et al. PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr. D 66, 213–221 (2010).

  53. 53.

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

  54. 54.

    Goddard, T. D. et al. UCSF ChimeraX: meeting modern challenges in visualization and analysis. Protein Sci. 27, 14–25 (2018).

  55. 55.

    Waterhouse, A. M., Procter, J. B., Martin, D. M., Clamp, M. & Barton, G. J. Jalview version 2—a multiple sequence alignment editor and analysis workbench. Bioinformatics 25, 1189–1191 (2009).

  56. 56.

    Liu, F. & Heck, A. J. Interrogating the architecture of protein assemblies and protein interaction networks by cross-linking mass spectrometry. Curr. Opin. Struct. Biol. 35, 100–108 (2015).

  57. 57.

    Liu, F., Lössl, P., Scheltema, R., Viner, R. & Heck, A. J. R. Optimized fragmentation schemes and data analysis strategies for proteome-wide cross-link identification. Nat. Commun. 8, 15473 (2017).

  58. 58.

    Grimm, M., Zimniak, T., Kahraman, A. & Herzog, F. xVis: a web server for the schematic visualization and interpretation of crosslink-derived spatial restraints. Nucleic Acids Res. 43, W362–W369 (2015).

  59. 59.

    Vizcaíno, J. A. et al. ProteomeXchange provides globally coordinated proteomics data submission and dissemination. Nat. Biotechnol. 32, 223–226 (2014).

  60. 60.

    Yan, K., Zhang, Z., Yang, J., McLaughlin, S. H. & Barford, D. Architecture of the CBF3–centromere complex of the budding yeast kinetochore. Nat. Struct. Mol. Biol. 25, 1103–1110 (2018).

  61. 61.

    Hinshaw, S. M., Dates, A. N. & Harrison, S. C. The structure of the yeast Ctf3 complex. eLife 8, e48215 (2019).

  62. 62.

    Akiyoshi, B. et al. Tension directly stabilizes reconstituted kinetochore-microtubule attachments. Nature 468, 576–579 (2010).

  63. 63.

    Gonen, S. et al. The structure of purified kinetochores reveals multiple microtubule-attachment sites. Nat. Struct. Mol. Biol. 19, 925–929 (2012).

Download references

Acknowledgements

This work was funded by MRC grant (MC_UP_1201/6) and CRUK grant (C576/A14109) to D.B., Horizon 2020 program INFRAIA project Epic-XS (Project 823839) to A.J.R.H. and Deutsche Forschungsgemeinschaft (EH237/12-1) to A.E.E.-M. We thank the LMB, eBIC and the Universities of Cambridge and Leeds Electron Microscopy facilities for help with the electron microscopy data collection, S. Scheres for help with electron microscopy processing, members of the Barford group for useful discussions, J. Grimmett and T. Darling for computing and J. Shi for help with insect cell expression.

Reviewer information

Nature thanks Eva Nogales and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

Author information

Z.Z. cloned kinetochore and nucleosome constructs. J.Y. and Z.Z. purified proteins, performed the protein-complex reconstitutions and biochemical and genetic analyses. K.Y. and L.C. prepared cryo-EM grids, collected and analysed electron microscopy data and determined the 3D reconstructions of CCAN−Cenp-ANuc and free Cenp-HIK, respectively. D.B. and K.Y. fitted coordinates and built models and J.Y. and S.H.M. performed SEC–MALS and analytical ultracentrifugation. D.F. collected and analysed XL-MS data. A.J.R.H. directed XL-MS experiments and analysis. A.E.E.-M. generated the chl4Δ cse4-R37A and chl4Δ yeast strains. D.B. directed the project. K.Y. and D.B. wrote the manuscript with help from all authors.

Correspondence to David Barford.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Extended data figures and tables

Extended Data Fig. 1 Reconstituted S. cerevisiae CCAN–Cenp-ANuc complexes.

a, Size-exclusion chromatogram profiles (Agilent Bio SEC-5 column) for (i) CCAN, (ii) CCAN–Cenp-A nucleosome (with Widom 601) complex, (iii) Cenp-A nucleosome (with Widom 601), (iv) H3 nucleosome (with Widom 601) and (v) H3N-Cenp-ANuc (with Widom 601). b, Comparative size-exclusion chromatogram profiles (Agilent Bio SEC-5 column) for CCAN–Cenp-ANuc with the Cenp-A nucleosome wrapped with either the (i) 147-bp Widom 601 positioning sequence (CCAN–Cenp-ANuc (Widom 601) as in a) or (ii) a 153-bp S. cerevisiae centromeric Cen3 sequence (CCAN–Cenp-ANuc (Cen3)). Both complexes eluted at the same volume. CCAN and the H3 nucleosome do not form a complex (iii). c, Coomassie-blue-stained SDS–PAGE of the 14-subunit CCAN complex. d, Coomassie-blue-stained SDS–PAGE gel of Cenp-ANuc (Widom 601). Lane E32, ethidium bromide-stained gel of fraction 32. e, CCAN–Cenp-ANuc (Widom 601) complex. Lane E13, ethidium-bromide-stained gel of fraction 13. Size-exclusion chromatograms are shown in a. f, SDS–PAGE gel of CCAN and H3 nucleosome (Widom 601) SEC run shown in b. gj, Coomassie-blue-stained SDS–PAGE gels of various Cenp-H, I and K segments co-expressed with Cenp-TW and purified with a double Strep tag on the tagged Cenp-I subunit (*). j, The HFDs of Cenp-TW (Cenp-THFDW) interact with the Cenp-HIKHead. These results confirm the assignments of the Cenp-H, K and I subunits in our cryo-EM maps. k, Schematic of the organization of CCAN–Cenp-ANuc subunits and subcomplexes and connections to the outer kinetochore Mis12 and Ndc80 complexes. Lines indicate subcomplex connections. The two pathways connecting Cenp-ANuc to the Ndc80 complex and microtubules are indicated as P1 and P2 (thick lines to Ndc80). Subunits of the essential P1 pathway are labelled black and indicated with blue shading, whereas subunits of the non-essential P2 pathway are labelled white and indicated with yellow shading. The P2 pathway becomes essential when the P1 pathway is defective through defects in Dsn1 phosphorylation9. The experiments shown in aj were performed independently in triplicate with similar results. For gel source data, see Supplementary Fig. 1.

Extended Data Fig. 2 Cryo-EM data of the S. cerevisiae CCAN–Cenp-ANuc complex.

a, A typical cryo-electron micrograph of CCAN–Cenp-ANuc, representative of 9,002 micrographs. b, Galleries of 2D classes of CCAN, representative of 100 2D classes. c, Galleries of 2D classes of CCAN–Cenp-ANuc, representative of 150 2D classes. The 2D class averages for the C2-symmetric (CCAN)2–Cenp-ANuc complex viewed in the plane of the C2-symmetry axis are outlined in red. Only a few views were observed, precluding a 3D reconstruction. Cryo-EM grids partially destabilize CCAN–Cenp-ANuc interactions, resulting in a very low abundance of (CCAN)2–Cenp-ANuc particles (about 0.03% of total). The two-fold symmetry axes of the (CCAN)2–Cenp-ANuc complex are shown as dashed arrows. Experiments for data in b and c were performed independently 12 times with similar results. d, FSC curves shown for the cryo-EM reconstructions of CCAN–Cenp-ANuc complexes: apo-CCAN, mask1 (Cenp-OPQU+, Cenp-LN), mask2 (Cenp-HIK, Cenp-LN, sub-Cenp-OP), CCAN–Cenp-ANuc. Mask1 and mask2 used for MBR are defined in h and i and Methods. e, Angular distribution plot of CCAN–Cenp-ANuc particles. f, Local resolution map of CCAN. g, Local resolution map of CCAN–Cenp-ANuc. h, Local resolution map of mask1 (Cenp-OPQU+, Cenp-LN). i, Local resolution map of mask2 (Cenp-HIK, Cenp-LN, sub-Cenp-OP).

Extended Data Fig. 3 Workflow of 3D classification of the CCAN–Cenp-ANuc cryo-EM dataset.

a, After initial 2D classification, about 1.4 million particles were sorted by 3D classification into apo-CCAN (52%) and the CCAN–Cenp-ANuc complex (48%). For apo-CCAN, 4% existed as dimers (black box) and 19% showed an ordered head-group (Cenp-HIKHead) for the Cenp-HIK–TW subcomplex (blue box). A mask was applied to the CCAN–Cenp-ANuc cryo-EM map to exclude the structurally variable Cenp-HIKHead domain for reconstruction of the 4.15 Å structure. b, Details of the four masks used for MBR. c, A small 3D class of CCAN–Cenp-ANuc, revealing density attached to Cenp-HIKHead contacting the DNA gyre of Cenp-ANuc was assigned as Cenp-THFDW.

Extended Data Fig. 4 Cryo-EM density maps of apo-CCAN.

a, Portion of cryo-EM map for the coiled coils of Cenp-H and Cenp-K. A selection of highly conserved intersubunit residues defined in b and c are labelled. These residues are well defined in the cryo-EM density, consistent with the structure. b, c, Multiple sequence alignment of the coiled-coil regions of Cenp-H (b) and Cenp-K (c). df, Portions of cryo-EM maps for Cenp-LN (d), Cenp-I (e) and Nkp1–Nkp2 (f). The chain assignments and polarity of Cenp-H, Cenp-I and Cenp-K of our structure agree with the cryo-EM structure of yeast Ctf3 (PDB: 6OUA)61.

Extended Data Fig. 5 Cryo-EM densities of CCAN and CCAN–Cenp-ANuc complexes.

a, Cryo-EM reconstruction of CCAN–Cenp-ANuc from uncrosslinked sample at 8.6 Å resolution. b, Cryo-EM map of dimeric CCAN (also Extended Data Fig. 3a, black box). Subunits are colour-coded as in Fig. 1. The 3.5 Å monomeric free CCAN coordinates were rigid-body-docked into the cryo-EM map. c, Cartoon representation of the S. cerevisiae MIND complex15 (right), showing a notable similarity to the coiled coils of Cenp-QU–Nkp1–Nkp2 of CENP-OPQU+ (left). d, View of the 4.7 Å resolution cryo-EM map of free Cenp-HIK with fitted coordinates from CCAN. e, In the context of CCAN, Cenp-HIKHead rotates to accommodate Cenp-ANuc. The two conformations of Cenp-HIK from the apo-CCAN and CCAN–Cenp-ANuc complexes were superimposed onto their rigid portion of Cenp-HIK (C-terminal region of Cenp-I is shown for apo-CCAN) to indicate the conformational variability of Cenp-HIKHead between the two states. Subunits of Cenp-HIKHead of CCAN–Cenp-ANuc are coloured lighter. f, Cryo-EM density of Cenp-ANuc showing the Cenp-C motif of Cenp-C.

Extended Data Fig. 6 XL-MS analysis of the CCAN and CCAN–Cenp-ANuc complexes.

a, b, Circular plots displaying all the identified crosslinks for CCAN (a) and CCAN–Cenp-ANuc (b). Inter- and intra-subunit crosslinks are indicated in red and blue, respectively c, d, Histogram plots showing the Cα–Cα distance distribution of the crosslinks that could be mapped onto the CCAN (c) and CCAN–Cenp-ANuc structures (d). Ninety-five per cent of the mapped crosslinks satisfy the crosslinker-imposed distance restraint of 30 Å indicated with a dashed red line. e, f, Crosslinks mapped onto the CCAN (e) and CCAN–Cenp-ANuc complex (f). Inter and intra-subunit crosslinks are indicated in red and blue, respectively. Crosslinks exceeding the crosslinker-imposed distance restraint of 30 Å are indicated in yellow. g, Residues on CCAN shown by XL-MS that crosslink with Cenp-C are indicated on the CCAN structure. Red spheres, crosslinks in the CCAN–Cenp-ANuc complex; yellow spheres, additional crosslinks unique to apo-CCAN. The experiments shown in a and b were performed independently in triplicate with similar results.

Extended Data Fig. 7 The S. cerevisiae Cenp-ANuc nucleosome is unwrapped.

ac, The positively charged electrostatic potential of the DNA-binding groove of Cenp-LN subcomplex is conserved in S. cerevisiae, S. pombe and H. sapiens. S. pombe and H. sapiens are represented by modelled structures. d, Cenp-N interacts with S. cerevisiae Cenp-ANuc in the context of CCAN differently from the interaction of free human Cenp-N with Cenp-ANuc. The Cenp-N subunit of the human Cenp-N–Cenp-A nucleosome structure (PDB: 6C0W29) was superimposed onto Cenp-N of the S. cerevisiae CCAN–Cenp-ANuc structure. In this mode of Cenp-N–Cenp-ANuc interactions, Cenp-ANuc would clash with Cenp-OPQU+ and Cenp-N of CCAN. e, f, Structures of S. cerevisiae H3Nuc (PDB: 1ID324) (e) and Cenp-ANuc (f, this work). g, Sequence alignment of the N-terminal regions of S. cerevisiae H3 and Cenp-A (Cse4) histones. For the chimeric H3N–Cenp-ANuc, residues 1–50 of S. cerevisiae H3 were substituted for residues 1–140 of S. cerevisiae Cenp-A. A similar approach was used for vertebrate Cenp-ANuc (ref. 23).

Extended Data Fig. 8 SDS–PAGE of CCANΔCenp-C–Cenp-ANuc complexes.

Corresponding size-exclusion chromatograms are shown in Fig. 4b and Extended Data Fig. 9a. a, b, Mutating the Cenp-N DNA-binding groove did not impair CCANΔCenp-C assembly. c, Wild-type CCANΔCenp-C forms a complex with Cenp-ANuc. d, Mutating the Cenp-N DNA-binding groove disrupts CCANΔCenp-C–Cenp-ANuc interactions. e, Mutating the L1 loop of Cenp-A did not destabilize CCANΔCenp-C–Cenp-ANuc interactions. f, Deletion of the N terminus of Cenp-A (1–129) (ΔNCenp-ANuc) did not impair CCANΔCenp-C–Cenp-ANuc interactions. h, Both CCANΔCenp-C and CCANΔCenp-C–Cenp-NMut bound poorly to H3N–Cenp-ANuc. The experiments shown were performed independently in triplicate with similar results. For gel source data, see Supplementary Fig. 1.

Extended Data Fig. 9 Testing of CCANΔCenp-C binding to Cenp-ANuc.

a, Comparative SEC profiles (Agilent Bio SEC-5 column) for wild-type CCANΔCenp-C and the Cenp-NMut of CCANΔCenp-C to Cenp-ANuc and its modifications (Cenp-ANuc-L1Nuc, ΔNCenp-ANuc and H3N-Cenp-ANuc) and H3Nuc. Mutating the L1 loop (Cenp-AL1-Nuc) of Cenp-A or deletion of the N-terminal 129 residues (ΔNCenp-ANuc) did not destabilize CCANΔCenp-C–Cenp-ANuc interactions. By contrast, CCAN with the Cenp-NMut bound less well and both CCAN and CCAN–Cenp-NMut hardly bound to H3N–Cenp-ANuc. (CCANΔC, CCANΔCenp-C). Associated SDS–PAGE is shown in Extended Data Figs. 8, 9b). b, Coomassie-blue-stained SDS–PAGE showed that CCANΔCenp-C did not associate with H3Nuc. c, Micrococcal nuclease digestion of Cenp-ANuc, H3Nuc and H3N–Cenp-ANuc. Widom 601 DNA is shown as a control. The H3Nuc and H3N–Cenp-ANuc protect a similar and longer length of DNA compared with Cenp-ANuc. d, Model of CBF360 bound to CCAN–Cenp-ANuc, indicating that CBF3 would not associate with a fully assembled kinetochore, consistent with proteomic data62. The experiments shown in ac were performed independently in triplicate with similar results. For gel source data, see Supplementary Fig. 1.

Extended Data Fig. 10 S. cerevisiae CCAN–Cenp-ANuc comprises two CCAN complexes in solution.

ac, The predicted mass of (CCAN)2–Cenp-ANuc is 1.31 MDa, (CCAN)1–Cenp-ANuc is 0.77 MDa and that of a CCAN dimer 1.09 MDa (Extended Data Table 2). Representative SEC–MALS data for crosslinked S. cerevisiae CCAN–Cenp-ANuc complex (a), run independently in triplicate with similar results, average molecular mass is 1.23 MDa ((CCAN)2–Cenp-ANuc); uncrosslinked S. cerevisiae CCAN–Cenp-ANuc complex (b), run independently in triplicate with similar results, with average masses of 1.38 MDa ((CCAN)2–Cenp-ANuc) and 526 kDa (CCAN); and S. cerevisiae CCAN alone (c), run independently in duplicate with similar results, with average masses of 839 kDa for the leading edge (green) and 650 kDa for the trailing edge (magenta), suggesting a non-resolved monomer–dimer equilibrium. d, e, Velocity analytical ultracentrifugation of crosslinked (d) and uncrosslinked (e) S. cerevisiae CCAN–Cenp-ANuc complexes with residuals to the fits shown in f and g. f, g, Fit of a c(s) distribution model for the crosslinked complex (f), the major species sediments at 15.8S (Sw,20 = 26.1S) with a minor species at 12.1S (Sw,20 = 20.0S) that corresponds to calculated masses of 1.34 MDa ((CCAN)2–Cenp-ANuc) and 896 kDa (possibly (CCAN)1–Cenp-ANuc), respectively, with a fitted value of 1.761 for the frictional ratio. g, Fit for uncrosslinked samples, the major species is resolved into two species that sediment at 14.3S (Sw,20 = 22.6S) and 15.7S (Sw,20 = 24.9S) with a minor species at 12.3S (Sw,20 = 19.4 S), which gave masses of 1.32 MDa ((CCAN)2–Cenp-ANuc) and 1.15 MDa ((CCAN)2) for the major species and 716 kDa ((CCAN)1–Cenp-ANuc) for the minor species. The experiments shown in dg were performed independently in triplicate with similar results. h, Examples of two 2D class averages showing the (CCAN)2–Cenp-ANuc particles viewed in the plane of the C2 symmetry axis (red outline) (data from Extended Data Fig. 2c) and the 2D reprojections of a modelled (CCAN)2–Cenp-ANuc based on the CCAN–Cenp-ANuc cryo-EM reconstruction (yellow outline) (shown in i). There is a close correspondence in shape and dimensions between the calculated reprojections and the observed 2D classes. The two-fold symmetry axes of the (CCAN)2–Cenp-ANuc complex are shown as dashed arrows. i, j, Two alternative models for how CCAN assembled onto a Cenp-A nucleosome would interact with the outer kinetochore–microtubule interface (Supplementary Video 2). i, In scenario (1), CCAN interacts with the outer kinetochore from the same side as the DNA-binding surface. Microtubules attached to the outer kinetochore would hoist CCAN from below the over-lying nucleosome and out-stretched DNA. j, In scenario (2), the microtubule-outer kinetochore interface contacts CCAN from the opposite side to the CCAN DNA-binding surface. Outer-kinetochore (outer-KT): KMN network and microtubule-attachment complexes, Dam1–DASH (budding yeast) and Ska proteins of vertebrates. The combined dimension of (CCAN)2–Cenp-ANuc (32 nm) matches that of the hub at the centre of the yeast kinetochore63.

Extended Data Table 1 Cryo-EM data collection, refinement and validation statistics
Extended Data Table 2 Table of CCAN subunits

Supplementary information

Supplementary Figure 1

Original source images for all data obtained by electrophoretic separation: Coomassie blue stained and ethidium bromide stained SDS PAGE and western blots and ethidium bromide stained agarose gel

Reporting Summary

Supplementary Table 1

Cross-linking mass spectrometry data for apo CCAN. Listed are cross-linked lysine residues

Supplementary Table 2

Cross-linking mass spectrometry data for CCAN–Cenp-ANuc. Listed are cross-linked lysine residues

Video 1

Cryo-EM density map for CCAN–Cenp-ANuc is shown, first without density for Cenp-HIKHead. Next, density map for CCAN–Cenp-ANuc with Cenp-HIKHead is shown

Video 2

Model for dimeric CCAN assembled onto Cenp-ANuc. The two scenarios for how CCAN–Cenp-ANuc might attach to the outer kinetochore are indicated (Extended Data Fig. 10i, j)

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Yan, K., Yang, J., Zhang, Z. et al. Structure of the inner kinetochore CCAN complex assembled onto a centromeric nucleosome. Nature 574, 278–282 (2019) doi:10.1038/s41586-019-1609-1

Download citation

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.