Three-dimensional genome architecture: players and mechanisms

  • A Corrigendum to this article was published on 05 August 2015

Key Points

  • Contacts between distant genomic regions in the same or in different chromosomes are important in the regulation of gene expression, as highlighted by the activation of genes by chromatin contacts between promoters and enhancers that can lie hundreds of kb away.

  • Chromatin contacts are currently measured by two main approaches: chromosome conformation capture (3C)-based techniques and nuclear imaging methods such as fluorescence in situ hybridization (FISH). Both approaches have caveats, and the field is ripe for further technical development.

  • The formation of chromatin contacts is promoted by chromatin-binding proteins that can bind two or more genomic regions simultaneously. Such proteins include transcription factors, RNA and DNA polymerases, Polycomb repressive complexes and chromosomal scaffold proteins such as cohesin.

  • Topologically associating domains (TADs) are genomic regions enriched with contacts within them. TADs have specific sizes and positions in the genome and are found in a wide range of metazoans. The factors and mechanisms that promote TAD formation are a matter of considerable debate.

  • The three-dimensional organization of the genome also depends on the formation of chromatin contacts with nuclear domains and compartments such as the nuclear lamina and the nucleolus. Specific sets of chromatin contacts are formed within each chromosome, and between them and nuclear domains. The mechanisms that govern chromosome localization, volume and shape remain poorly understood.

  • Many cellular processes such as division, differentiation and senescence, present challenges to the maintenance of nuclear organization, gene expression programs and cell identity. At the same time, they can also offer opportunities for chromatin remodelling and the reinforcement of gene expression patterns.


The different cell types of an organism share the same DNA, but during cell differentiation their genomes undergo diverse structural and organizational changes that affect gene expression and other cellular functions. These can range from large-scale folding of whole chromosomes or of smaller genomic regions, to the re-organization of local interactions between enhancers and promoters, mediated by the binding of transcription factors and chromatin looping. The higher-order organization of chromatin is also influenced by the specificity of the contacts that it makes with nuclear structures such as the lamina. Sophisticated methods for mapping chromatin contacts are generating genome-wide data that provide deep insights into the formation of chromatin interactions, and into their roles in the organization and function of the eukaryotic cell nucleus.

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Figure 1: Chromosome organization.
Figure 2: Topologically associating domains.
Figure 3: The nuclear envelope affects genome organization and function.
Figure 4: Chromatin structure of post-mitotic cells.

Change history

  • 05 August 2015

    In the original article, the following sentence was incorrect: “Histone marks associated with enhancers, such as histone 3 Lys4 monomethylation (H3K4me1), and with transcription repression, such as histone 3 Lys9 trimethylation (H3K9me3), were also enriched at TAD boundaries.2” The corrected sentence is as follows: “Histone marks associated with enhancers, such as histone 3 Lys4 monomethylation (H3K4me1), and with transcription repression, such as histone 3 Lys9 trimethylation (H3K9me3), were not found to be enriched at TAD boundaries.” This has been corrected in the online version of the article.


  1. 1

    Wijgerde, M., Grosveld, F. & Fraser, P. Transcription complex stability and chromatin dynamics in vivo. Nature 377, 209–213 (1995).

    CAS  PubMed  PubMed Central  Google Scholar 

  2. 2

    Dillon, N., Trimborn, T., Strouboulis, J., Fraser, P. & Grosveld, F. The effect of distance on long-range chromatin interactions. Mol. Cell 1, 131–139 (1997).

    CAS  PubMed  Google Scholar 

  3. 3

    Dekker, J., Rippe, K., Dekker, M. & Kleckner, N. Capturing chromosome conformation. Science 295, 1306–1311 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  4. 4

    Simonis, M. et al. Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C). Nature Genet. 38, 1348–1354 (2006).

    CAS  PubMed  Google Scholar 

  5. 5

    Naka, K. & Hirao, A. Maintenance of genomic integrity in hematopoietic stem cells. Int. J. Hematol. 93, 434–439 (2011).

    CAS  PubMed  Google Scholar 

  6. 6

    Stadhouders, R. et al. Multiplexed chromosome conformation capture sequencing for rapid genome-scale high-resolution detection of long-range chromatin interactions. Nature Protoc. 8, 509–524 (2013).

    CAS  Google Scholar 

  7. 7

    Dostie, J. et al. Chromosome Conformation Capture Carbon Copy (5C): a massively parallel solution for mapping interactions between genomic elements. Genome Res. 16, 1299–1309 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  8. 8

    Hughes, J. R. et al. Analysis of hundreds of cis-regulatory landscapes at high resolution in a single, high-throughput experiment. Nature Genet. 46, 205–212 (2014).

    CAS  PubMed  Google Scholar 

  9. 9

    Rodley, C. D., Bertels, F., Jones, B. & O'Sullivan, J. M. Global identification of yeast chromosome interactions using genome conformation capture. Fungal Genet. Biol. 46, 879–886 (2009).

    CAS  PubMed  Google Scholar 

  10. 10

    Lieberman-Aiden, E. et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009).

    CAS  Article  Google Scholar 

  11. 11

    Kalhor, R., Tjong, H., Jayathilaka, N., Alber, F. & Chen, L. Genome architectures revealed by tethered chromosome conformation capture and population-based modeling. Nature Biotech. 30, 90–98 (2012).

    CAS  Google Scholar 

  12. 12

    Kolovos, P. et al. Targeted Chromatin Capture (T2C): a novel high resolution high throughput method to detect genomic interactions and regulatory elements. Epigenet. Chromatin 7, 10 (2014).

    Google Scholar 

  13. 13

    Tolhuis, B., Palstra, R. J., Splinter, E., Grosveld, F. & de Laat, W. Looping and interaction between hypersensitive sites in the active β-globin locus. Mol. Cell 10, 1453–1465 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  14. 14

    Vernimmen, D., De Gobbi, M., Sloane-Stanley, J. A., Wood, W. G. & Higgs, D. R. Long-range chromosomal interactions regulate the timing of the transition between poised and active gene expression. EMBO J. 26, 2041–2051 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  15. 15

    Stadhouders, R. et al. Dynamic long-range chromatin interactions control Myb proto-oncogene transcription during erythroid development. EMBO J. 31, 986–999 (2012).

    CAS  PubMed  Google Scholar 

  16. 16

    Jing, H. et al. Exchange of GATA factors mediates transitions in looped chromatin organization at a developmentally regulated gene locus. Mol. Cell 29, 232–242 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  17. 17

    Markova, E. N., Kantidze, O. L. & Razin, S. V. Transcriptional regulation and spatial organisation of the human AML1/RUNX1 gene. J. Cell Biochem. 112, 1997–2005 (2011).

    CAS  PubMed  Google Scholar 

  18. 18

    Blackledge, N. P., Ott, C. J., Gillen, A. E. & Harris, A. An insulator element 3′ to the CFTR gene binds CTCF and reveals an active chromatin hub in primary cells. Nucleic Acids Res. 37, 1086–1094 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. 19

    Ktistaki, E. et al. CD8 locus nuclear dynamics during thymocyte development. J. Immunol. 184, 5686–5695 (2010).

    CAS  PubMed  Google Scholar 

  20. 20

    Palstra, R. J. et al. The β-globin nuclear compartment in development and erythroid differentiation. Nature Genet. 35, 190–194 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  21. 21

    Love, P. E., Warzecha, C. & Li, L. Ldb1 complexes: the new master regulators of erythroid gene transcription. Trends Genet. 30, 1–9 (2014).

    CAS  PubMed  Google Scholar 

  22. 22

    Deng, W. et al. Controlling long-range genomic interactions at a native locus by targeted tethering of a looping factor. Cell 149, 1233–1244 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  23. 23

    Andersson, R. et al. An atlas of active enhancers across human cell types and tissues. Nature 507, 455–461 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  24. 24

    Grosveld, F., van Assendelft, G. B., Greaves, D. R. & Kollias, G. Position-independent, high-level expression of the human β-globin gene in transgenic mice. Cell 51, 975–985 (1987).

    CAS  PubMed  Google Scholar 

  25. 25

    Sabbattini, P., Georgiou, A., Sinclair, C. & Dillon, N. Analysis of mice with single copies and multiple copies of transgenes reveals a novel arrangement for the λ5-VpreB1 locus control region. Mol. Cell. Biol. 19, 671–679 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  26. 26

    Fields, P. E., Lee, G. R., Kim, S. T., Bartsevich, V. V. & Flavell, R. A. Th2-specific chromatin remodeling and enhancer activity in the Th2 cytokine locus control region. Immunity 21, 865–876 (2004).

    CAS  PubMed  Google Scholar 

  27. 27

    Ellis, J., Talbot, D., Dillon, N. & Grosveld, F. Synthetic human β-globin 5′HS2 constructs function as locus control regions only in multicopy transgene concatamers. EMBO J. 12, 127–134 (1993).

    CAS  PubMed  PubMed Central  Google Scholar 

  28. 28

    Nasmyth, K. & Haering, C. H. Cohesin: its roles and mechanisms. Annu. Rev. Genet. 43, 525–558 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. 29

    Parelho, V. et al. Cohesins functionally associate with CTCF on mammalian chromosome arms. Cell 132, 422–433 (2008).

    CAS  Google Scholar 

  30. 30

    Hadjur, S. et al. Cohesins form chromosomal cis-interactions at the developmentally regulated IFNG locus. Nature 460, 410–413 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  31. 31

    Seitan, V. C. et al. A role for cohesin in T-cell-receptor rearrangement and thymocyte differentiation. Nature 476, 467–471 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  32. 32

    Lobanenkov, V. V. et al. A novel sequence-specific DNA binding protein which interacts with three regularly spaced direct repeats of the CCCTC-motif in the 5′-flanking sequence of the chicken c-myc gene. Oncogene 5, 1743–1753 (1990).

    CAS  PubMed  Google Scholar 

  33. 33

    Ong, C. T. & Corces, V. G. CTCF: an architectural protein bridging genome topology and function. Nature Rev. Genet. 15, 234–246 (2014).

    CAS  PubMed  Google Scholar 

  34. 34

    Sanyal, A., Lajoie, B. R., Jain, G. & Dekker, J. The long-range interaction landscape of gene promoters. Nature 489, 109–113 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  35. 35

    Xu, Z., Wei, G., Chepelev, I., Zhao, K. & Felsenfeld, G. Mapping of INS promoter interactions reveals its role in long-range regulation of SYT8 transcription. Nature Struct. Mol. Biol. 18, 372–378 (2011).

    CAS  Google Scholar 

  36. 36

    Kehayova, P., Monahan, K., Chen, W. & Maniatis, T. Regulatory elements required for the activation and repression of the protocadherin-α gene cluster. Proc. Natl Acad. Sci. USA 108, 17195–17200 (2011).

    CAS  PubMed  Google Scholar 

  37. 37

    Guo, Y. et al. CTCF/cohesin-mediated DNA looping is required for protocadherin-α promoter choice. Proc. Natl Acad. Sci. USA 109, 21081–21086 (2012).

    CAS  PubMed  Google Scholar 

  38. 38

    Dixon, J. R. et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485, 376–380 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  39. 39

    Nora, E. P. et al. Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 485, 381–385 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  40. 40

    Hou, C., Li, L., Qin, Z. S. & Corces, V. G. Gene density, transcription, and insulators contribute to the partition of the Drosophila genome into physical domains. Mol. Cell 48, 471–484 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. 41

    Sexton, T. et al. Three-dimensional folding and functional organization principles of the Drosophila genome. Cell 148, 458–472 (2012).

    CAS  Google Scholar 

  42. 42

    Van Bortle, K. et al. Insulator function and topological domain border strength scale with architectural protein occupancy. Genome Biol. 15, R82 (2014).

    PubMed  PubMed Central  Google Scholar 

  43. 43

    Kim, Y. J., Cecchini, K. R. & Kim, T. H. Conserved, developmentally regulated mechanism couples chromosomal looping and heterochromatin barrier activity at the homeobox gene A locus. Proc. Natl Acad. Sci. USA 108, 7391–7396 (2011).

    CAS  PubMed  Google Scholar 

  44. 44

    Seitan, V. C. et al. Cohesin-based chromatin interactions enable regulated gene expression within preexisting architectural compartments. Genome Res. 23, 2066–2077 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. 45

    Zuin, J. et al. Cohesin and CTCF differentially affect chromatin architecture and gene expression in human cells. Proc. Natl Acad. Sci. USA 111, 996–1001 (2014).

    CAS  Google Scholar 

  46. 46

    Sofueva, S. et al. Cohesin-mediated interactions organize chromosomal domain architecture. EMBO J. 32, 3119–3129 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. 47

    Young, S. G., Jung, H. J., Coffinier, C. & Fong, L. G. Understanding the roles of nuclear A- and B-type lamins in brain development. J. Biol. Chem. 287, 16103–16110 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  48. 48

    Houben, F. et al. Disturbed nuclear orientation and cellular migration in A-type lamin deficient cells. Biochim. Biophys. Acta 1793, 312–324 (2009).

    CAS  PubMed  Google Scholar 

  49. 49

    Dechat, T., Adam, S. A., Taimen, P., Shimi, T. & Goldman, R. D. Nuclear lamins. CSH Persp. Biol. 2, a000547 (2010).

    CAS  Google Scholar 

  50. 50

    Amendola, M. & van Steensel, B. Mechanisms and dynamics of nuclear lamina-genome interactions. Curr. Opin. Cell Biol. 28, 61–68 (2014).

    CAS  PubMed  Google Scholar 

  51. 51

    Guelen, L. et al. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453, 948–951 (2008).

    CAS  PubMed  Google Scholar 

  52. 52

    Meuleman, W. et al. Constitutive nuclear lamina–genome interactions are highly conserved and associated with A/T-rich sequence. Genome Res. 23, 270–280 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  53. 53

    Finlan, L. E. et al. Recruitment to the nuclear periphery can alter expression of genes in human cells. PLoS Genet. 4, e1000039 (2008).

    PubMed  PubMed Central  Google Scholar 

  54. 54

    Lundgren, M. et al. Transcription factor dosage affects changes in higher order chromatin structure associated with activation of a heterochromatic gene. Cell 103, 733–743 (2000).

    CAS  PubMed  Google Scholar 

  55. 55

    Kind, J. et al. Single-cell dynamics of genome–nuclear lamina interactions. Cell 153, 178–192 (2013).

    CAS  PubMed  Google Scholar 

  56. 56

    Padeken, J. & Heun, P. Nucleolus and nuclear periphery: velcro for heterochromatin. Curr. Opin. Cell Biol. 28, 54–60 (2014).

    CAS  PubMed  Google Scholar 

  57. 57

    van Koningsbruggen, S. et al. High-resolution whole-genome sequencing reveals that specific chromatin domains from most human chromosomes associate with nucleoli. Mol. Biol. Cell 21, 3735–3748 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  58. 58

    Nemeth, A. et al. Initial genomics of the human nucleolus. PLoS Genet. 6, e1000889 (2010).

    PubMed  PubMed Central  Google Scholar 

  59. 59

    Parada, L. A., McQueen, P. G. & Misteli, T. Tissue-specific spatial organization of genomes. Genome Biol. 5, R44 (2004).

    PubMed  PubMed Central  Google Scholar 

  60. 60

    Tanabe, H. et al. Evolutionary conservation of chromosome territory arrangements in cell nuclei from higher primates. Proc. Natl Acad. Sci. USA 99, 4424–4429 (2002).

    CAS  PubMed  Google Scholar 

  61. 61

    Bridger, J. M. Chromobility: the rapid movement of chromosomes in interphase nuclei. Biochem. Soc. Trans. 39, 1747–1751 (2011).

    CAS  PubMed  Google Scholar 

  62. 62

    Williams, R. R., Broad, S., Sheer, D. & Ragoussis, J. Subchromosomal positioning of the epidermal differentiation complex (EDC) in keratinocyte and lymphoblast interphase nuclei. Exp. Cell Res. 272, 163–175 (2002).

    CAS  PubMed  Google Scholar 

  63. 63

    Ferrai, C. et al. Poised transcription factories prime silent uPA gene prior to activation. PLoS Biol. 8, e1000270 (2010).

    PubMed  PubMed Central  Google Scholar 

  64. 64

    Volpi, E. et al. Large-scale chromatin organisation of the major histocompatibility complex and other regions of human chromosome 6 and its response to interferon in interphase nuclei J. Cell Sci. 113, 1565–1576 (2000).

    CAS  PubMed  Google Scholar 

  65. 65

    Meaburn, K. J., Gudla, P. R., Khan, S., Lockett, S. J. & Misteli, T. Disease-specific gene repositioning in breast cancer. J. Cell Biol. 187, 801–812 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  66. 66

    Cremer, T. et al. Chromosome territories — a functional nuclear landscape. Curr. Opin. Cell Biol. 18, 307–316 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  67. 67

    Branco, M. R. & Pombo, A. Intermingling of chromosome territories in interphase suggests role in translocations and transcription-dependent associations. PLoS Biol. 4, e138 (2006).

    PubMed  PubMed Central  Google Scholar 

  68. 68

    Branco, M. R., Branco, T., Ramirez, F. & Pombo, A. Changes in chromosome organization during PHA-activation of resting human lymphocytes measured by cryo-FISH. Chromosome Res. 16, 413–426 (2008).

    CAS  PubMed  Google Scholar 

  69. 69

    Zhang, Y. et al. Spatial organization of the mouse genome and its role in recurrent chromosomal translocations. Cell 148, 908–921 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  70. 70

    Hakim, O. et al. DNA damage defines sites of recurrent chromosomal translocations in B lymphocytes. Nature 484, 69–74 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  71. 71

    Roukos, V. et al. Spatial dynamics of chromosome translocations in living cells. Science 341, 660–664 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  72. 72

    Jackson, D. A. & Pombo, A. Replicon clusters are stable units of chromosome structure: evidence that nuclear organization contributes to the efficient activation and propagation of S phase in human cells. J. Cell Biol. 140, 1285–1295 (1998).

    CAS  PubMed  PubMed Central  Google Scholar 

  73. 73

    Ma, H. et al. Spatial and temporal dynamics of DNA replication sites in mammalian cells. J. Cell Biol. 143, 1415–1425 (1998).

    CAS  PubMed  PubMed Central  Google Scholar 

  74. 74

    Gottesfeld, J. M. & Forbes, D. J. Mitotic repression of the transcriptional machinery. Trends Biochem. Sci. 22, 197–202 (1997).

    CAS  PubMed  Google Scholar 

  75. 75

    Belmont, A. S. Mitotic chromosome structure and condensation. Curr. Opin. Cell Biol. 18, 632–638 (2006).

    CAS  PubMed  Google Scholar 

  76. 76

    Naumova, N. et al. Organization of the mitotic chromosome. Science 342, 948–953 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  77. 77

    Chow, C.-M. et al. Variant histone H3.3 marks promoters of transcriptionally active genes during mammalian cell division. EMBO Rep. 6, 354–360 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  78. 78

    Kouskouti, A. & Talianidis, I. Histone modifications defining active genes persist after transcriptional and mitotic inactivation. EMBO J. 24, 347–357 (2004).

    PubMed  PubMed Central  Google Scholar 

  79. 79

    Kelly, T. K. et al. H2A.Z maintenance during mitosis reveals nucleosome shifting on mitotically silenced genes. Mol. Cell 39, 901–911 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  80. 80

    Caravaca, J. M. et al. Bookmarking by specific and nonspecific binding of FoxA1 pioneer factor to mitotic chromosomes. Genes Dev. 27, 251–260 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  81. 81

    Kadauke, S. et al. Tissue-specific mitotic bookmarking by hematopoietic transcription factor GATA1. Cell 150, 725–737 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  82. 82

    Young, D. W. et al. Mitotic retention of gene expression patterns by the cell fate-determining transcription factor Runx2. Proc. Natl Acad. Sci. USA 104, 3189–3194 (2007).

    CAS  Google Scholar 

  83. 83

    Blobel, G. A. et al. A reconfigured pattern of MLL occupancy within mitotic chromatin promotes rapid transcriptional reactivation following mitotic exit. Mol. Cell 36, 970–983 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  84. 84

    Zhao, R., Nakamura, T., Fu, Y., Lazar, Z. & Spector, D. L. Gene bookmarking accelerates the kinetics of post-mitotic transcriptional re-activation. Nature Cell Biol. 13, 1295–1304 (2011).

    CAS  PubMed  Google Scholar 

  85. 85

    Rawlings, J. S., Gatzka, M., Thomas, P. G. & Ihle, J. N. Chromatin condensation via the condensin II complex is required for peripheral T-cell quiescence. EMBO J. 30, 263–276 (2011).

    CAS  PubMed  Google Scholar 

  86. 86

    Sabbattini, P. et al. An H3K9/S10 methyl-phospho switch modulates Polycomb and Pol II binding at repressed genes during differentiation. Mol. Biol. Cell 25, 904–915 (2014).

    PubMed  PubMed Central  Google Scholar 

  87. 87

    Frangini, A. et al. The Aurora B kinase and the polycomb protein Ring1B combine to regulate active promoters in quiescent lymphocytes. Mol. Cell 51, 647–661 (2013).

    CAS  PubMed  Google Scholar 

  88. 88

    Sabbattini, P. et al. A novel role for the Aurora B kinase in epigenetic marking of silent chromatin in differentiated postmitotic cells. EMBO J. 26, 4657–4669 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  89. 89

    Fischle, W. et al. Regulation of HP1-chromatin binding by histone H3 methylation and phosphorylation. Nature 438, 1116–1122 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  90. 90

    Solovei, I. et al. LBR and lamin A/C sequentially tether peripheral heterochromatin and inversely regulate differentiation. Cell 152, 584–598 (2013).

    CAS  PubMed  Google Scholar 

  91. 91

    Solovei, I. et al. Nuclear architecture of rod photoreceptor cells adapts to vision in mammalian evolution. Cell 137, 356–368 (2009).

    CAS  PubMed  Google Scholar 

  92. 92

    Helmlinger, D. et al. Glutamine-expanded ataxin-7 alters TFTC/STAGA recruitment and chromatin structure leading to photoreceptor dysfunction. PLoS Biol. 4, e67 (2006).

    PubMed  PubMed Central  Google Scholar 

  93. 93

    Rai, T. S. & Adams, P. D. Lessons from senescence: chromatin maintenance in non-proliferating cells. Biochim. Biophys. Acta 1819, 322–331 (2013).

    PubMed  Google Scholar 

  94. 94

    Zhang, R., Chen, W. & Adams, P. D. Molecular dissection of formation of senescence-associated heterochromatin foci. Mol. Cell. Biol. 27, 2343–2358 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  95. 95

    Ye, X. et al. Downregulation of Wnt signaling is a trigger for formation of facultative heterochromatin and onset of cell senescence in primary human cells. Mol. Cell 27, 183–196 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  96. 96

    Freund, A., Laberge, R. M., Demaria, M. & Campisi, J. Lamin B1 loss is a senescence-associated biomarker. Mol. Biol. Cell 23, 2066–2075 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  97. 97

    Shimi, T. et al. The role of nuclear lamin B1 in cell proliferation and senescence. Genes Dev. 25, 2579–2593 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  98. 98

    Shah, P. P. et al. Lamin B1 depletion in senescent cells triggers large-scale changes in gene expression and the chromatin landscape. Genes Dev. 27, 1787–1799 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  99. 99

    Sadaie, M. et al. Redistribution of the lamin B1 genomic binding profile affects rearrangement of heterochromatic domains and SAHF formation during senescence. Genes Dev. 27, 1800–1808 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  100. 100

    Friedl, P., Wolf, K. & Lammerding, J. Nuclear mechanics during cell migration. Curr. Opin. Cell Biol. 23, 55–64 (2011).

    CAS  PubMed  Google Scholar 

  101. 101

    Mohrin, M. et al. Hematopoietic stem cell quiescence promotes error-prone DNA repair and mutagenesis. Cell Stem Cell 7, 174–185 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  102. 102

    Rehen, S. K. et al. Chromosomal variation in neurons of the developing and adult mammalian nervous system. Proc. Natl Acad. Sci. USA 98, 13361–13366 (2001).

    CAS  Google Scholar 

  103. 103

    Duncan, A. W. et al. The ploidy conveyor of mature hepatocytes as a source of genetic variation. Nature 467, 707–710 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  104. 104

    Terns, R. M. & Terns, M. P. CRISPR-based technologies: prokaryotic defense weapons repurposed. Trends Genet. 30, 111–118 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. 105

    Dekker, J., Marti-Renom, M. A. & Mirny, L. A. Exploring the three-dimensional organization of genomes: interpreting chromatin interaction data. Nature Rev. Genet. 14, 390–403 (2013).

    CAS  Google Scholar 

  106. 106

    Ethier, S. D., Miura, H. & Dostie, J. Discovering genome regulation with 3C and 3C-related technologies. Biochim. Biophys. Acta 1819, 401–410 (2012).

    CAS  Google Scholar 

  107. 107

    de Wit, E. & de Laat, W. A decade of 3C technologies: insights into nuclear organization. Genes Dev. 26, 11–24 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  108. 108

    van de Werken, H. J. et al. Robust 4C-seq data analysis to screen for regulatory DNA interactions. Nature Methods 9, 969–972 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  109. 109

    Belmont, A. S. Large-scale chromatin organization: the good, the surprising, and the still perplexing. Curr. Opin. Cell Biol. 26, 69–78 (2014).

    CAS  PubMed  Google Scholar 

  110. 110

    Gavrilov, A. A. et al. Disclosure of a structural milieu for the proximity ligation reveals the elusive nature of an active chromatin hub. Nucleic Acids Res. 41, 3563–3575 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  111. 111

    O'Sullivan, J. M. Hendy, M. D., Pichugina, T., Wake, G. C. & Langowski, J. The statistical-mechanics of chromosome conformation capture. Nucleus 4, 390–398 (2013).

    PubMed  PubMed Central  Google Scholar 

  112. 112

    Nicodemi, M. & Pombo, A. Models of chromosome structure. Curr. Opin. Cell Biol. 28, 90–95 (2014).

    CAS  PubMed  Google Scholar 

  113. 113

    Barbieri, M. et al. Complexity of chromatin folding is captured by the strings and binders switch model. Proc. Natl Acad. Sci. USA 109, 16173–16178 (2012).

    CAS  PubMed  Google Scholar 

  114. 114

    Bau, D. et al. The three-dimensional folding of the α-globin gene domain reveals formation of chromatin globules. Nature Struct. Mol. Biol. 18, 107–114 (2011).

    CAS  Google Scholar 

  115. 115

    Nagano, T. et al. Single-cell Hi-C reveals cell-to-cell variability in chromosome structure. Nature 502, 59–64 (2013).

    CAS  Google Scholar 

  116. 116

    Hozak, P. & Cook, P. R. Replication factories. Trends Cell Biol. 4, 48–52 (1994).

    CAS  PubMed  Google Scholar 

  117. 117

    Baddeley, D. et al. Measurement of replication structures at the nanometer scale using super-resolution light microscopy. Nucleic Acids Res. 38, e8 (2010).

    CAS  PubMed  Google Scholar 

  118. 118

    Jackson, D. A., Iborra, F. J., Manders, E. M. & Cook, P. R. Numbers and organization of RNA polymerases, nascent transcripts, and transcription units in HeLa nuclei. Mol. Biol. Cell 9, 1523–1536 (1998).

    CAS  PubMed  PubMed Central  Google Scholar 

  119. 119

    Martin, S. & Pombo, A. Transcription factories: quantitative studies of nanostructures in the mammalian nucleus. Chromosome Res. 11, 461–470 (2003).

    CAS  PubMed  Google Scholar 

  120. 120

    Pombo, A. et al. Specialized transcription factories within mammalian nuclei. Crit. Rev. Eukaryot. Gene Expr 10, 21–29 (2000).

    CAS  PubMed  Google Scholar 

  121. 121

    Kimura, H., Tao, Y., Roeder, R. G. & Cook, P. R. Quantitation of RNA polymerase II and its transcription factors in an HeLa cell: little soluble holoenzyme but significant amounts of polymerases attached to the nuclear substructure. Mol. Cell. Biol. 19, 5383–5392 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  122. 122

    Pombo, A. et al. Regional specialization in human nuclei: visualization of discrete sites of transcription by RNA polymerase III. EMBO J. 18, 2241–2253 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  123. 123

    Faro-Trindade, I. & Cook, P. R. A conserved organization of transcription during embryonic stem cell differentiation and in cells with high C value. Mol. Biol. Cell 17, 2910–2920 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  124. 124

    Jackson, D. A. Features of nuclear architecture that influence gene expression in higher eukaryotes: confronting the enigma of epigenetics. J. Cell Biochem. 79(Suppl.35), 69–77 (2000).

    Google Scholar 

  125. 125

    Osborne, C. et al. Active genes dynamically colocalize to shared sites of ongoing transcription. Nature Genet. 36, 1065–1071 (2004).

    CAS  PubMed  Google Scholar 

  126. 126

    Schoenfelder, S. et al. Preferential associations between co-regulated genes reveal a transcriptional interactome in erythroid cells. Nature Genet. 42, 53–61 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  127. 127

    Brookes, E. et al. Polycomb associates genome-wide with a specific RNA polymerase II variant, and regulates metabolic genes in ESCs. Cell Stem Cell 10, 157–170 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  128. 128

    Lanzuolo, C., Roure, V., Dekker, J., Bantignies, F. & Orlando, V. Polycomb response elements mediate the formation of chromosome higher-order structures in the bithorax complex. Nature Cell Biol. 9, 1167–1174 (2007).

    CAS  PubMed  Google Scholar 

  129. 129

    Grimaud, C. et al. RNAi components are required for nuclear clustering of Polycomb group response elements. Cell 124, 957–971 (2006).

    CAS  PubMed  Google Scholar 

  130. 130

    Tiwari, V. K., Cope, L., McGarvey, K. M., Ohm, J. E. & Baylin, S. B. A novel 6C assay uncovers Polycomb-mediated higher order chromatin conformations. Genome Res. 18, 1171–1179 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

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The authors thank P. Sabbattini for providing the images shown in Fig. 4a. Work in N.D.'s laboratory is supported by the Medical Research Council, UK. A.P. thanks the Helmholtz Foundation for support.

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Correspondence to Ana Pombo or Niall Dillon.

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The authors declare no competing financial interests.

Supplementary information

Supplementary information S1 (box)

Relating chromatin contacts to gene regulation mechanisms (PDF 117 kb)

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Chromatin immunoprecipitation

A method in which chromatin bound by a protein is immunoprecipitated with an antibody against that protein, to allow the extraction and analysis of the bound DNA by quantitative PCR or genome-wide sequencing.

Nuclear lamina

A protein meshwork made of intermediate filaments (such as lamins) and membrane-associated proteins (such as emerin) that covers the inner nuclear membrane and is responsible for maintaining nuclear shape, organization and function.


Highly condensed chromatin that shows dark staining. Constitutive heterochromatin remains in this state throughout the cell cycle. Facultative heterochromatin is cell-type-specific condensed chromatin that is often a feature of terminally differentiated cells.

DNA adenine methyltransferase identification

A method based on expression of fusion proteins with bacterial Dam methylase, and detection of methylated DNA as a measure of its contact with the fusion protein.

Cryoprotected cells

Cells that have been treated with a cryoproctectant to prevent structural damage during freezing.

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Pombo, A., Dillon, N. Three-dimensional genome architecture: players and mechanisms. Nat Rev Mol Cell Biol 16, 245–257 (2015).

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