Coupling 1D modifications and 3D nuclear organization: data, models and function

Current Opinion in Cell Biology

Volumn 44, Issue , 2017, Pages 20-27

  Jost, Daniel ^a   Vaillant, Cédric ^b   Meister, Peter ^c  

^a UNIV GRENOBLE ALPES   (France)

^b UNIVERSITÉ LYON 1   (France)

^c UNIVERSITY OF BERN   (Switzerland)

Author keywords

[No Author keywords available]

Indexed keywords

POLYMER; CHROMATIN; DNA;

CELL COMPARTMENTALIZATION; CELL NUCLEAR STRUCTURE; CHROMATIN; CHROMOSOME; CONFORMATION; DNA MODIFICATION; GENE CONTROL; GENETIC LINKAGE; GENETIC ORGANIZATION; HUMAN; MICROSCOPY; NONHUMAN; PRIORITY JOURNAL; REVIEW; ANIMAL; CELL NUCLEUS; CHEMISTRY; GENE EXPRESSION REGULATION; GENOME; MOLECULAR MODEL;

ANIMALS; CELL NUCLEUS; CHROMATIN; DNA; GENE EXPRESSION REGULATION; GENOME; HUMANS; MODELS, MOLECULAR;

EID: 85007356745     PISSN: 09550674     EISSN: 18790410     Source Type: Journal    
DOI: 10.1016/j.ceb.2016.12.001     Document Type: Review

References (80)

1. 1 Dekker, J., Rippe, K., Dekker, M., Kleckner, N., Capturing chromosome conformation. Science 295 (2002), 1306–1311.
2. 2 de Wit, E., de Laat, W., A decade of 3C technologies: insights into nuclear organization. Genes Dev 26 (2012), 11–24.
3. 3 Nora, E.P., Lajoie, B.R., Schulz, E.G., Giorgetti, L., Okamoto, I., Servant, N., Piolot, T., van Berkum, N.L., Meisig, J., Sedat, J., et al. Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 485 (2012), 381–385.
4. 4•• Rao, S.S., Huntley, M.H., Durand, N.C., Stamenova, E.K., Bochkov, I.D., Robinson, J.T., Sanborn, A.L., Machol, I., Omer, A.D., Lander, E.S., et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159 (2014), 1665–1680 The authors perform high resolution Hi-C in different cell lines and different experimental conditions (fixed/unfixed). Beyond the TAD structure of the genome, they identify loops between convergent CTCF binding sites, which are conserved between the mouse and the human genome.
5. 5• Boettiger, A.N., Bintu, B., Moffitt, J.R., Wang, S., Beliveau, B.J., Fudenberg, G., Imakaev, M., Mirny, L.A., Wu, C.-t., Zhuang, X., Super-resolution imaging reveals distinct chromatin folding for different epigenetic states. Nature 529 (2016), 418–422 Using fluorescent in situ hybridization and 3D Stochastic Optical Reconstruction Microscopy (STORM), a super resolution technique, the authors demonstrate that Polycomb, active and unmodified histone H3 epigenomic domains have different compaction levels. Moreover, they observe directly that epigenetically similar domains preferentially contact each other.
6. 6 Fabre, P.J., Benke, A., Joye, E., Nguyen Huynh, T.H., Manley, S., Duboule, D., Nanoscale spatial organization of the HoxD gene cluster in distinct transcriptional states. Proc Natl Acad Sci U S A 112 (2015), 13964–13969.
7. 7 Lieberman-Aiden, E., van Berkum, N.L., Williams, L., Imakaev, M., Ragoczy, T., Telling, A., Amit, I., Lajoie, B.R., Sabo, P.J., Dorschner, M.O., et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326 (2009), 289–293.
8. 8 Marbouty, M., Cournac, A., Flot, J.F., Marie-Nelly, H., Mozziconacci, J., Koszul, R., Metagenomic chromosome conformation capture (meta3C) unveils the diversity of chromosome organization in microorganisms. Elife, 3, 2014, e03318.
9. 9 Sexton, T., Yaffe, E., Kenigsberg, E., Bantignies, F., Leblanc, B., Hoichman, M., Parrinello, H., Tanay, A., Cavalli, G., Three-dimensional folding and functional organization principles of the Drosophila genome. Cell 148 (2012), 458–472.
10. 10 Dixon, J.R., Selvaraj, S., Yue, F., Kim, A., Li, Y., Shen, Y., Hu, M., Liu, J.S., Ren, B., Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485 (2012), 376–380.
11. 11 Frenster, J.H., Allfrey, V.G., Mirsky, A.E., Repressed and active chromatin isolated from interphase lymphocytes. Proc Natl Acad Sci U S A 50 (1963), 1026–1032.
12. 12 Dekker, J., Heard, E., Structural and functional diversity of topologically associating domains. FEBS Lett 589 (2015), 2877–2884.
13. 13• Ghavi-Helm, Y., Klein, F.A., Pakozdi, T., Ciglar, L., Noordermeer, D., Huber, W., Furlong, E.E., Enhancer loops appear stable during development and are associated with paused polymerase. Nature 512 (2014), 96–100 By focusing on enhancer/promoter contacts during two stages of Drosophila development and two different tissues, the authors observe that promoter/enhancer contacts are mainly unchanged between transcriptionally inactive and active states. This suggests that enhancer/promoter contacts are preformed and structurally organize the genome.
14. 14 Guo, Y., Xu, Q., Canzio, D., Shou, J., Li, J., Gorkin, D.U., Jung, I., Wu, H., Zhai, Y., Tang, Y., et al. CRISPR inversion of CTCF sites alters genome topology and enhancer/promoter function. Cell 162 (2015), 900–910.
15. 15•• Lupianez, D.G., Kraft, K., Heinrich, V., Krawitz, P., Brancati, F., Klopocki, E., Horn, D., Kayserili, H., Opitz, J.M., Laxova, R., et al. Disruptions of topological chromatin domains cause pathogenic rewiring of gene-enhancer interactions. Cell 161 (2015), 1012–1025 The authors of this study demonstrate that disruption of a TAD boundary in both mice and humans leads to ectopic contacts between a promoter and an enhancer located in a nearby TAD. This ultimately leads to misregulation of the Epha4 gene and finger malformations.
16. 16 Franke, M., Ibrahim, D.M., Andrey, G., Schwarzer, W., Heinrich, V., Schopflin, R., Kraft, K., Kempfer, R., Jerkovic, I., Chan, W.L., et al. Formation of new chromatin domains determines pathogenicity of genomic duplications. Nature 538 (2016), 265–269.
17. 17•• Dixon, J.R., Jung, I., Selvaraj, S., Shen, Y., Antosiewicz-Bourget, J.E., Lee, A.Y., Ye, Z., Kim, A., Rajagopal, N., Xie, W., et al. Chromatin architecture reorganization during stem cell differentiation. Nature 518 (2015), 331–336 This paper describes TAD structures in human ES cells and 4 derived lineages. The TAD structure itself is remarkable stable between cell types, while the contact frequencies inside TADs and between TADs varies, affecting 36% of the genome.
18. 18 Lonfat, N., Montavon, T., Darbellay, F., Gitto, S., Duboule, D., Convergent evolution of complex regulatory landscapes and pleiotropy at Hox loci. Science 346 (2014), 1004–1006.
19. 19 Jin, F., Li, Y., Dixon, J.R., Selvaraj, S., Ye, Z., Lee, A.Y., Yen, C.A., Schmitt, A.D., Espinoza, C.A., Ren, B., A high-resolution map of the three-dimensional chromatin interactome in human cells. Nature 503 (2013), 290–294.
20. 20 Le Dily, F., Bau, D., Pohl, A., Vicent, G.P., Serra, F., Soronellas, D., Castellano, G., Wright, R.H., Ballare, C., Filion, G., et al. Distinct structural transitions of chromatin topological domains correlate with coordinated hormone-induced gene regulation. Genes Dev 28 (2014), 2151–2162.
21. 21 Wong, H., Marie-Nelly, H., Herbert, S., Carrivain, P., Blanc, H., Koszul, R., Fabre, E., Zimmer, C., A predictive computational model of the dynamic 3D interphase yeast nucleus. Curr Biol 22 (2012), 1881–1890.
22. 22 Kimura, H., Shimooka, Y., Nishikawa, J.-i., Miura, O., Sugiyama, S., Yamada, S., Ohyama, T., The genome folding mechanism in yeast. J Biochem 154 (2013), 137–147.
23. 23 Avşaroğlu Bi, Bronk, G., Gordon-Messer, S., Ham, J., Bressan, D.A., Haber, J.E., Kondev, J., Effect of chromosome tethering on nuclear organization in yeast. PLoS One, 9, 2014, e102474.
24. 24 Rosa, A., Everaers, R., Structure and dynamics of interphase chromosomes. PLoS Comput Biol, 4, 2008, e1000153.
25. 25 Bohn, M., Heermann, D.W., Diffusion-driven looping provides a consistent framework for chromatin organization. PLoS One, 5, 2010, e12218.
26. 26 Halverson, J.D., Smrek, J., Kremer, K., Grosberg, A.Y., From a melt of rings to chromosome territories: the role of topological constraints in genome folding. Rep Prog Phys, 77, 2014, 022601.
27. 27 Gürsoy, G., Xu, Y., Kenter, A.L., Liang, J., Spatial confinement is a major determinant of the folding landscape of human chromosomes. Nucleic Acids Res 42 (2014), 8223–8230.
28. 28 Grosberg, A., Rabin, Y., Halvin, S., Neer, A., The crumpled globule model of the three-dimensional structure of DNA. Eur Phys Lett, 23, 1993, 373.
29. 29 Mirny, L.A., The fractal globule as a model of chromatin architecture in the cell. Chromosome Res 19 (2011), 37–51.
30. 30 Ho, J.W., Jung, Y.L., Liu, T., Alver, B.H., Lee, S., Ikegami, K., Sohn, K.A., Minoda, A., Tolstorukov, M.Y., Appert, A., et al. Comparative analysis of metazoan chromatin organization. Nature 512 (2014), 449–452.
31. 31 Zhu, Y., Chen, Z., Zhang, K., Wang, M., Medovoy, D., Whitaker, J.W., Ding, B., Li, N., Zheng, L., Wang, W., Constructing 3D interaction maps from 1D epigenomes. Nat Commun, 7, 2016, 10812.
32. 32 Fraser, J., Ferrai, C., Chiariello, A.M., Schueler, M., Rito, T., Laudanno, G., Barbieri, M., Moore, B.L., Kraemer, D.C., Aitken, S., et al. Hierarchical folding and reorganization of chromosomes are linked to transcriptional changes in cellular differentiation. Mol Syst Biol, 11, 2015, 852.
33. 33 Barbieri, M., Chotalia, M., Fraser, J., Lavitas, L.-M., Dostie, J., Pombo, A., Nicodemi, M., Complexity of chromatin folding is captured by the strings and binders switch model. Proc Natl Acad Sci U S A 109 (2012), 16173–16178.
34. 34 Brackley, C.A., Johnson, J., Kelly, S., Cook, P.R., Marenduzzo, D., Simulated binding of transcription factors to active and inactive regions folds human chromosomes into loops, rosettes and topological domains. Nucleic Acids Res 44 (2016), 3503–3512.
35. 35 Jerabek, H., Heermann, D.W., Expression-dependent folding of interphase chromatin. PLoS One, 7, 2012, e37525.
36. 36 Benedetti, F., Dorier, J., Burnier, Y., Stasiak, A., Models that include supercoiling of topological domains reproduce several known features of interphase chromosomes. Nucleic Acids Res 42 (2014), 2848–2855.
37. 37 Doyle, B., Fudenberg, G., Imakaev, M., Mirny, L.A., Chromatin loops as allosteric modulators of enhancer-promoter interactions. PLoS Comput Biol, 10, 2014, e1003867.
38. 38 Ganai, N., Sengupta, S., Menon, G.I., Chromosome positioning from activity-based segregation. Nucleic Acids Res 42 (2014), 4145–4159.
39. 39•• Jost, D., Carrivain, P., Cavalli, G., Vaillant, C., Modeling epigenome folding: formation and dynamics of topologically associated chromatin domains. Nucleic Acids Res 42 (2014), 9553–9561 A heteropolymer model is introduced which suggests that epigenomic-driven interactions might be an important driver of chromosome folding at megabase scales. In agreement with experimental data, the model predicts that TAD formation is fast and results from the internal collapse of epigenomic domain and that inter-TAD interactions are dynamic and emerge at longer time-scales.
40. 40 Tark-Dame, M., Jerabek, H., Manders, E.M.M., Heermann, D.W., van Driel, R., Depletion of the chromatin looping proteins CTCF and cohesin causes chromatin compaction: insight into chromatin folding by polymer modelling. PLoS Comput Biol, 10, 2014, e1003877.
41. 41 Nazarov, L.I., Tamm, M.V., Avetosov, V.A., Nechaev, S.K., A statistical model of intra-chromosome contact maps. Soft Matter 11 (2015), 1019–1025.
42. 42•• Sanborn, A.L., Rao, S.S., Huang, S.C., Durand, N.C., Huntley, M.H., Jewett, A.I., Bochkov, I.D., Chinnappan, D., Cutkosky, A., Li, J., et al. Chromatin extrusion explains key features of loop and domain formation in wild-type and engineered genomes. Proc Natl Acad Sci U S A 112 (2015), E6456–E6465 A polymer model for chromatin extrusion is introduced and shown to agree quantitatively with experimental data. The role of oriented CTCF sites is explored experimentally, showing that the model is able to predict the effect of CTCF site inversions or deletions.
43. 43 Ulianov, S.V., Khrameeva, E.E., Gavrilov, A.A., Flyamer, I.M., Kos, P., Mikhaleva, E.A., Penin, A.A., Logacheva, M.D., Imakaev, M.V., Chertovich, A., et al. Active chromatin and transcription play a key role in chromosome partitioning into topologically associating domains. Genome Res 26 (2016), 70–84.
44. 44•• Fudenberg, G., Imakaev, M., Lu, C., Goloborodko, A., Abdennur, N., Mirny, L.A., Formation of chromosomal domains by loop extrusion. Cell Rep 15 (2016), 2038–2049 This paper introduces a rigorous analysis of the loop extrusion model and the role of boundary elements [note that preprint publication of this work was first released in bioRxiv in August 2015]. Compared to other possible models, the loop extrusion model is shown to fit the data best. Polymer simulations and genomic data are used to identify the functional role of CTCF and cohesin.
45. 45 Tiana, G., Amitai, A., Pollex, T., Piolot, T., Holcman, D., Heard, E., Giorgetti, L., Structural fluctuations of the chromatin fiber within topologically associating domains. Biophys J 110 (2016), 1234–1245.
46. 46 Giorgetti, L., Galupa, R., Nora, E.P., Piolot, T., Lam, F., Dekker, J., Tiana, G., Heard, E., Predictive polymer modeling reveals coupled fluctuations in chromosome conformation and transcription. Cell 157 (2014), 950–963.
47. 47 Olarte-Plata, J.D., Haddad, N., Vaillant, C., Jost, D., The folding landscape of the epigenome. Phys Biol, 13, 2016, 026001.
48. 48 Care, B.R., Emeriau, P.-E., Cortini, R., Victor, J.-M., Chromatin epigenomic domain folding: size matters. AIMS Biophys 2 (2015), 517–530.
49. 49 Canzio, D., Liao, M., Naber, N., Pate, E., Larson, A., Wu, S., Marina, D.B., Garcia, J.F., Madhani, H.D., Cooke, R., et al. A conformational switch in HP1 releases auto-inhibition to drive heterochromatin assembly. Nature 496 (2013), 377–381.
50. 50 Isono, K., Endo, T.A., Ku, M., Yamada, D., Suzuki, R., Sharif, J., Ishikura, T., Toyoda, T., Bernstein, B.E., Koseki, H., SAM domain polymerization links subnuclear clustering of PRC1 to gene silencing. Dev Cell 26 (2013), 565–577.
51. 51 Hiragami-Hamada, K., Soeroes, S., Nikolov, M., Wilkins, B., Kreuz, S., Chen, C., De La Rosa-Velázquez, I.A., Zenn, H.M., Kost, N., Pohl, W., et al. Dynamic and flexible H3K9me3 bridging via HP1β dimerization establishes a plastic state of condensed chromatin. Nat Commun, 7, 2016, 11310.
52. 52• Wani, A.H., Boettiger, A.N., Schorderet, P., Ergun, A., Münger, C., Sadreyev, R.I., Zhuang, X., Kingston, R.E., Francis, N.J., Chromatin topology is coupled to Polycomb group protein subnuclear organization. Nat Commun, 7, 2016, 10291 This study makes use of a polymer model using interacting binders, first introduced by Barbieri et al. [33], to mechanistically dissect the formation of Polycomb bodies in flies. Results are then compared to in vivo superresolution microscopy data.
53. 53 Brackley, C.A., Brown, J.M., Waithe, D., Babbs, C., Davies, J., Hughes, J.R., Buckle, V.J., Marenduzzo, D., Predicting the three-dimensional folding of cis-regulatory regions in mammalian genomes using bioinformatic data and polymer models. Genome Biol, 17, 2016, 59.
54. 54 Naumova, N., Imakaev, M., Fudenberg, G., Zhan, Y., Lajoie, B.R., Mirny, L.A., Dekker, J., Organization of the mitotic chromosome. Science 342 (2013), 948–953.
55. 55 Nagano, T., Lubling, Y., Stevens, T.J., Schoenfelder, S., Yaffe, E., Dean, W., Laue, E.D., Tanay, A., Fraser, P., Single-cell Hi-C reveals cell-to-cell variability in chromosome structure. Nature 502 (2013), 59–64.
56. 56 Vietri Rudan, M., Barrington, C., Henderson, S., Ernst, C., Odom, D.T., Tanay, A., Hadjur, S., Comparative Hi-C reveals that CTCF underlies evolution of chromosomal domain architecture. Cell Rep 10 (2015), 1297–1309.
57. 57•• Dowen, J.M., Fan, Z.P., Hnisz, D., Ren, G., Abraham, B.J., Zhang, L.N., Weintraub, A.S., Schuijers, J., Lee, T.I., Zhao, K., et al. Control of cell identity genes occurs in insulated neighborhoods in mammalian chromosomes. Cell 159 (2014), 374–387 From a genome-wide SMC1 ChIA-PET experiment performed in murine ESC, the authors identify for the first time cohesin-mediated, TAD-like interaction domains with CTCF-enriched boundaries. By specific deletion of these boundaries they show how these domains, by constituting structural “insulated neighborhood” may contribute to the control of the superenhancer-mediated activity of pluripotent genes as well as the PcG-repression of lineage-specific genes.
58. 58 Nasmyth, K., Disseminating the genome: joining, resolving, and separating sister chromatids during mitosis and meiosis. Annu Rev Genet 35 (2001), 673–745.
59. 59 Alipour, E., Marko, J.F., Self-organization of domain structures by DNA-loop-extruding enzymes. Nucleic Acids Res 40 (2012), 11202–11212.
60. 60 Crane, E., Bian, Q., McCord, R.P., Lajoie, B.R., Wheeler, B.S., Ralston, E.J., Uzawa, S., Dekker, J., Meyer, B.J., Condensin-driven remodelling of X chromosome topology during dosage compensation. Nature 523 (2015), 240–244.
61. 61 Goloborodko, A., Imakaev, M.V., Marko, J.F., Mirny, L., Compaction and segregation of sister chromatids via active loop extrusion. Elife, 2016, 5.
62. 62 Goloborodko, A., Marko, J.F., Mirny, L.A., Chromosome compaction by active loop extrusion. Biophys J 110 (2016), 2162–2168.
63. 63 Vilar, J.M., Leibler, S., DNA looping and physical constraints on transcription regulation. J Mol Biol 331 (2003), 981–989.
64. 64 Oehler, S., Alberti, S., Muller-Hill, B., Induction of the lac promoter in the absence of DNA loops and the stoichiometry of induction. Nucleic Acids Res 34 (2006), 606–612.
65. 65 Oehler, S., Amouyal, M., Kolkhof, P., von Wilcken-Bergmann, B., Muller-Hill, B., Quality and position of the three lac operators of E. coli define efficiency of repression. EMBO J 13 (1994), 3348–3355.
66. 66 Saiz, L., Vilar, J.M., DNA looping: the consequences and its control. Curr Opin Struct Biol 16 (2006), 344–350.
67. 67 Vilar, J.M., Saiz, L., Systems biophysics of gene expression. Biophys J 104 (2013), 2574–2585.
68. 68 Spitz, F., Gene regulation at a distance: from remote enhancers to 3D regulatory ensembles. Semin Cell Dev Biol 57 (2016), 57–67.
69. 69 Lanzuolo, C., Orlando, V., The function of the epigenome in cell reprogramming. Cell Mol Life Sci 64 (2007), 1043–1062.
70. 70 Bantignies, F., Roure, V., Comet, I., Leblanc, B., Schuettengruber, B., Bonnet, J., Tixier, V., Mas, A., Cavalli, G., Polycomb-dependent regulatory contacts between distant Hox loci in Drosophila. Cell 144 (2011), 214–226.
71. 71 Pueschel, R., Coraggio, F., Meister, P., From single genes to entire genomes: the search for a function of nuclear organization. Development 143 (2016), 910–923.
72. 72 Beisel, C., Paro, R., Silencing chromatin: comparing modes and mechanisms. Nat Rev Genet 12 (2011), 123–135.
73. 73 Zhang, T., Cooper, S., Brockdorff, N., The interplay of histone modifications - writers that read. EMBO Rep 16 (2015), 1467–1481.
74. 74 Chen, T., Dent, S.Y., Chromatin modifiers and remodellers: regulators of cellular differentiation. Nat Rev Genet 15 (2014), 93–106.
75. 75 Obersriebnig, M.J., Pallesen, E.M., Sneppen, K., Trusina, A., Thon, G., Nucleation and spreading of a heterochromatic domain in fission yeast. Nat Commun, 7, 2016, 11518.
76. 76 Angel, A., Song, J., Dean, C., Howard, M., A polycomb-based switch underlying quantitative epigenetic memory. Nature 476 (2011), 105–108.
77. 77 Dodd, I.B., Micheelsen, M.A., Sneppen, K., Thon, G., Theoretical analysis of epigenetic cell memory by nucleosome modification. Cell 129 (2007), 813–822.
78. 78 Jost, D., Bifurcation in epigenetics: implications in development, proliferation, and diseases. Phys Rev E Stat Nonlin Soft Matter Phys, 89, 2014, 010701.
79. 79 Soshnev, A.A., Josefowicz, S.Z., Allis, C.D., Greater than the sum of parts: complexity of the dynamic epigenome. Mol Cell 62 (2016), 681–694.
80. 80 Simon, J.A., Kingston, R.E., Occupying chromatin: polycomb mechanisms for getting to genomic targets, stopping transcriptional traffic, and staying put. Mol Cell 49 (2013), 808–824.