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The relationship between genome structure and function

A Publisher Correction to this article was published on 14 October 2021

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Abstract

Precise patterns of gene expression in metazoans are controlled by three classes of regulatory elements: promoters, enhancers and boundary elements. During differentiation and development, these elements form specific interactions in dynamic higher-order chromatin structures. However, the relationship between genome structure and its function in gene regulation is not completely understood. Here we review recent progress in this field and discuss whether genome structure plays an instructive role in regulating gene expression or is a reflection of the activity of the regulatory elements of the genome.

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Fig. 1: The regulatory elements of the genome.
Fig. 2: The organization of the genome.
Fig. 3: Compartmentalization and loop extrusion.
Fig. 4: The relationship between the linear order of the regulatory elements and the organization of the genome.
Fig. 5: Formation of enhancer–promoter interactions by loop extrusion and affinity.

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References

  1. Cavalli, G. & Heard, E. Advances in epigenetics link genetics to the environment and disease. Nature 571, 489–499 (2019).

    CAS  PubMed  Google Scholar 

  2. Pang, B. & Snyder, M. P. Systematic identification of silencers in human cells. Nat. Genet. 52, 254–263 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  3. Ngan, C. Y. et al. Chromatin interaction analyses elucidate the roles of PRC2-bound silencers in mouse development. Nat. Genet. 52, 264–272 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  4. Kellis, M. et al. Defining functional DNA elements in the human genome. Proc. Natl Acad. Sci. USA 111, 6131 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  5. Dunham, I. et al. An integrated encyclopedia of DNA elements in the human genome. Nature 489, 57–74 (2012).

    CAS  Google Scholar 

  6. Furlong, E. E. M. & Levine, M. Developmental enhancers and chromosome topology. Science 361, 1341–1345 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  7. Schoenfelder, S. & Fraser, P. Long-range enhancer–promoter contacts in gene expression control. Nat. Rev. Genet. 20, 437–455 (2019).

    CAS  PubMed  Google Scholar 

  8. Rowley, M. J. & Corces, V. G. Organizational principles of 3D genome architecture. Nat. Rev. Genet. 19, 789–800 (2018).

    CAS  PubMed  Google Scholar 

  9. Stadhouders, R., Filion, G. J. & Graf, T. Transcription factors and 3D genome conformation in cell-fate decisions. Nature 569, 345–354 (2019).

    CAS  PubMed  Google Scholar 

  10. Kempfer, R. & Pombo, A. Methods for mapping 3D chromosome architecture. Nat. Rev. Genet. 21, 207–226 (2020).

    CAS  PubMed  Google Scholar 

  11. McCord, R. P., Kaplan, N. & Giorgetti, L. Chromosome conformation capture and beyond: toward an integrative view of chromosome structure and function. Mol. Cell 77, 688–708 (2020).

    CAS  PubMed  Google Scholar 

  12. Mirny, L. A., Imakaev, M. & Abdennur, N. Two major mechanisms of chromosome organization. Curr. Opin. Cell Biol. 58, 142–152 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. Robson, M. I., Ringel, A. R. & Mundlos, S. Regulatory landscaping: how enhancer-promoter communication is sculpted in 3D. Mol. Cell 74, 1110–1122 (2019).

    CAS  PubMed  Google Scholar 

  14. Janssen, A., Colmenares, S. U. & Karpen, G. H. Heterochromatin: guardian of the genome. Annu. Rev. Cell Dev. Biol. 34, 265–288 (2018).

    CAS  PubMed  Google Scholar 

  15. Berthelot, C., Villar, D., Horvath, J. E., Odom, D. T. & Flicek, P. Complexity and conservation of regulatory landscapes underlie evolutionary resilience of mammalian gene expression. Nat. Ecol. Evol. 2, 152–163 (2018).

    PubMed  Google Scholar 

  16. Andersson, R. & Sandelin, A. Determinants of enhancer and promoter activities of regulatory elements. Nat. Rev. Genet. 21, 71–87 (2020).

    CAS  PubMed  Google Scholar 

  17. Smale, S. T. & Kadonaga, J. T. The RNA polymerase II core promoter. Annu. Rev. Biochem. 72, 449–479 (2003).

    CAS  PubMed  Google Scholar 

  18. Baumann, M., Pontiller, J. & Ernst, W. Structure and basal transcription complex of RNA polymerase II core promoters in the mammalian genome: an overview. Mol. Biotechnol. 45, 241–247 (2010).

    CAS  PubMed  Google Scholar 

  19. Sainsbury, S., Bernecky, C. & Cramer, P. Structural basis of transcription initiation by RNA polymerase II. Nat. Rev. Mol. Cell Biol. 16, 129–143 (2015).

    CAS  PubMed  Google Scholar 

  20. Cramer, P. Organization and regulation of gene transcription. Nature 573, 45–54 (2019).

    CAS  PubMed  Google Scholar 

  21. Core, L. J., Waterfall, J. J. & Lis, J. T. Nascent RNA sequencing reveals widespread pausing and divergent initiation at human promoters. Science 322, 1845 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  22. Seila, A. C. et al. Divergent transcription from active promoters. Science 322, 1849 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  23. Preker, P. et al. RNA exosome depletion reveals transcription upstream of active human promoters. Science 322, 1851 (2008).

    CAS  PubMed  Google Scholar 

  24. Smale, S. T. Core promoters: active contributors to combinatorial gene regulation. Genes Dev. 15, 2503–2508 (2001).

    CAS  PubMed  Google Scholar 

  25. Cooper, S. J., Trinklein, N. D., Anton, E. D., Nguyen, L. & Myers, R. M. Comprehensive analysis of transcriptional promoter structure and function in 1% of the human genome. Genome Res. 16, 1–10 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  26. Su, W., Jackson, S., Tjian, R. & Echols, H. DNA looping between sites for transcriptional activation: self-association of DNA-bound Sp1. Genes Dev. 5, 820–826 (1991).

    CAS  PubMed  Google Scholar 

  27. Calhoun, V. C., Stathopoulos, A. & Levine, M. Promoter-proximal tethering elements regulate enhancer-promoter specificity in the Drosophila Antennapedia complex. Proc. Natl Acad. Sci. USA 99, 9243 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  28. Zabidi, M. A. et al. Enhancer–core-promoter specificity separates developmental and housekeeping gene regulation. Nature 518, 556–559 (2015).

    CAS  PubMed  Google Scholar 

  29. Banerji, J., Rusconi, S. & Schaffner, W. Expression of a beta-globin gene is enhanced by remote SV40 DNA sequences. Cell 27, 299–308 (1981).

    CAS  PubMed  Google Scholar 

  30. Long, H. K., Prescott, S. L. & Wysocka, J. Ever-changing landscapes: transcriptional enhancers in development and evolution. Cell 167, 1170–1187 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  31. Field, A. & Adelman, K. Evaluating enhancer function and transcription. Annu. Rev. Biochem. https://doi.org/10.1146/annurev-biochem-011420-095916 (2020).

  32. Hay, D. et al. Genetic dissection of the α-globin super-enhancer in vivo. Nat. Genet. 48, 895–903 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  33. Siersbæk, R. et al. Transcription factor cooperativity in early adipogenic hotspots and super-enhancers. Cell Rep. 7, 1443–1455 (2014).

    PubMed  Google Scholar 

  34. Allen, B. L. & Taatjes, D. J. The mediator complex: a central integrator of transcription. Nat. Rev. Mol. Cell Biol. 16, 155–166 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  35. Soutourina, J. Transcription regulation by the mediator complex. Nat. Rev. Mol. Cell Biol. 19, 262–274 (2018).

    CAS  PubMed  Google Scholar 

  36. 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 

  37. Shen, Y. et al. A map of the cis-regulatory sequences in the mouse genome. Nature 488, 116–120 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  38. Anderson, E. & Hill, R. E. Long range regulation of the sonic hedgehog gene. Curr. Opin. Genet. Dev. 27, 54–59 (2014).

    CAS  PubMed  Google Scholar 

  39. Montavon, T. et al. A regulatory archipelago controls Hox genes transcription in digits. Cell 147, 1132–1145 (2011).

    CAS  PubMed  Google Scholar 

  40. Osterwalder, M. et al. Enhancer redundancy provides phenotypic robustness in mammalian development. Nature 554, 239–243 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. Perry, M. W., Boettiger, A. N., Bothma, J. P. & Levine, M. Shadow enhancers foster robustness of Drosophila gastrulation. Curr. Biol. 20, 1562–1567 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  42. Frankel, N. et al. Phenotypic robustness conferred by apparently redundant transcriptional enhancers. Nature 466, 490–493 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  43. Whyte, W. A. et al. Master transcription factors and mediator establish super-enhancers at key cell identity genes. Cell 153, 307–319 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  44. Oudelaar, A. M. et al. Single-allele chromatin interactions identify regulatory hubs in dynamic compartmentalized domains. Nat. Genet. 50, 1744–1751 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. Allahyar, A. et al. Enhancer hubs and loop collisions identified from single-allele topologies. Nat. Genet. 50, 1151–1160 (2018).

    CAS  PubMed  Google Scholar 

  46. Ing-Simmons, E. et al. Spatial enhancer clustering and regulation of enhancer-proximal genes by cohesin. Genome Res. 25, 504–513 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. Moorthy, S. D. et al. Enhancers and super-enhancers have an equivalent regulatory role in embryonic stem cells through regulation of single or multiple genes. Genome Res. 27, 246–258 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  48. Pott, S. & Lieb, J. D. What are super-enhancers? Nat. Genet. 47, 8–12 (2015).

    CAS  PubMed  Google Scholar 

  49. Henriques, T. et al. Widespread transcriptional pausing and elongation control at enhancers. Genes Dev. 32, 26–41 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  50. Mikhaylichenko, O. et al. The degree of enhancer or promoter activity is reflected by the levels and directionality of eRNA transcription. Genes Dev. 32, 42–57 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  51. Nguyen, T. A. et al. High-throughput functional comparison of promoter and enhancer activities. Genome Res. 26, 1023–1033 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  52. Almada, A. E., Wu, X., Kriz, A. J., Burge, C. B. & Sharp, P. A. Promoter directionality is controlled by U1 snRNP and polyadenylation signals. Nature 499, 360–363 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  53. Kowalczyk, M. S. et al. Intragenic enhancers act as alternative promoters. Mol. Cell 45, 447–458 (2012).

    CAS  PubMed  Google Scholar 

  54. Dao, L. T. M. et al. Genome-wide characterization of mammalian promoters with distal enhancer functions. Nat. Genet. 49, 1073–1081 (2017).

    CAS  PubMed  Google Scholar 

  55. Bell, A. C., West, A. G. & Felsenfeld, G. The protein CTCF is required for the enhancer blocking activity of vertebrate insulators. Cell 98, 387–396 (1999).

    CAS  PubMed  Google Scholar 

  56. Kellum, R. & Schedl, P. A position-effect assay for boundaries of higher order chromosomal domains. Cell 64, 941–950 (1991).

    CAS  PubMed  Google Scholar 

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

    CAS  PubMed  Google Scholar 

  58. Wendt, K. S. et al. Cohesin mediates transcriptional insulation by CCCTC-binding factor. Nature 451, 796–801 (2008).

    CAS  PubMed  Google Scholar 

  59. 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 

  60. Rao, S. S. P. et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159, 1665–1680 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  61. Merkenschlager, M. & Nora, E. P. CTCF and cohesin in genome folding and transcriptional gene regulation. Annu. Rev. Genomics Hum. Genet. 17, 17–43 (2016).

    CAS  PubMed  Google Scholar 

  62. Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944.e922 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  63. Huang, H. et al. CTCF mediates dosage and sequence-context-dependent transcriptional insulation through formation of local chromatin domains. Preprint at bioRxiv https://doi.org/10.1101/2020.07.07.192526 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  64. Kaaij, L. J. T., Mohn, F., van der Weide, R. H., de Wit, E. & Bühler, M. The ChAHP complex counteracts chromatin looping at CTCF sites that emerged from SINE expansions in mouse. Cell 178, 1437–1451.e1414 (2019).

    CAS  PubMed  Google Scholar 

  65. Hansen, A. S. et al. Distinct classes of chromatin loops revealed by deletion of an RNA-binding region in CTCF. Mol. Cell 76, 395–411.e313 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  66. Saldaña-Meyer, R. et al. RNA interactions are essential for CTCF-mediated genome organization. Mol. Cell 76, 412–422.e415 (2019).

    PubMed  PubMed Central  Google Scholar 

  67. Hsieh, T.-H. S. et al. Resolving the 3D landscape of transcription-linked mammalian chromatin folding. Mol. Cell 78, 539–553.e8 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  68. Krietenstein, N. et al. Ultrastructural details of mammalian chromosome architecture. Mol. Cell 78, 554–565.e7, e557 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  69. Rowley, M. J. et al. Evolutionarily conserved principles predict 3D chromatin organization. Mol. Cell 67, 837–852.e837 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  70. Harrold, C. L. et al. A functional overlap between actively transcribed genes and chromatin boundary elements. Preprint at bioRxiv https://doi.org/10.1101/2020.07.01.182089 (2020).

    Article  Google Scholar 

  71. De Gobbi, M. et al. A regulatory SNP causes a human genetic disease by creating a new transcriptional promoter. Science 312, 1215–1217 (2006).

    PubMed  Google Scholar 

  72. Cho, S. W. et al. Promoter of lncRNA Gene PVT1 Is a tumor-suppressor DNA boundary element. Cell 173, 1398–1412.e1322 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  73. Enver, T. et al. Developmental regulation of human fetal-to-adult globin gene switching in transgenic mice. Nature 344, 309–313 (1990).

    CAS  PubMed  Google Scholar 

  74. Oudelaar, A. M. et al. A revised model for promoter competition based on multi-way chromatin interactions at the α-globin locus. Nat. Commun. 10, 1–8 (2019).

    CAS  Google Scholar 

  75. Fukaya, T., Lim, B. & Levine, M. Enhancer control of transcriptional bursting. Cell 166, 358–368 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  76. Cremer, T. & Cremer, C. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat. Rev. Genet. 2, 292–301 (2001).

    CAS  PubMed  Google Scholar 

  77. Baptista, J. et al. Molecular cytogenetic analyses of breakpoints in apparently balanced reciprocal translocations carried by phenotypically normal individuals. Eur. J. Hum. Genet. 13, 1205–1212 (2005).

    CAS  PubMed  Google Scholar 

  78. Ghavi-Helm, Y. et al. Highly rearranged chromosomes reveal uncoupling between genome topology and gene expression. Nat. Genet. 51, 1272–1282 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  79. Bonev, B. & Cavalli, G. Organization and function of the 3D genome. Nat. Rev. Genet. 17, 772–772 (2016).

    CAS  PubMed  Google Scholar 

  80. Gibcus, J. H. & Dekker, J. The hierarchy of the 3D genome. Mol. Cell 49, 773–782 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  82. Bickmore, W. A. & van Steensel, B. Genome architecture: domain organization of interphase chromosomes. Cell 152, 1270–1284 (2013).

    CAS  PubMed  Google Scholar 

  83. Esposito, A. et al. Models of polymer physics for the architecture of the cell nucleus. WIREs Syst. Biol. Med. 11, e1444 (2019).

    Google Scholar 

  84. Hildebrand, E. M. & Dekker, J. Mechanisms and functions of chromosome compartmentalization. Trends Biochem. Sci. 45, 385–396 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. Feodorova, Y., Falk, M., Mirny, L. A. & Solovei, I. Viewing nuclear architecture through the eyes of nocturnal mammals. Trends Cell Biol. 30, 276–289 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  86. van Steensel, B. & Furlong, E. E. M. The role of transcription in shaping the spatial organization of the genome. Nat. Rev. Mol. Cell Biol. 20, 290 (2019).

    Google Scholar 

  87. Rada-Iglesias, A., Grosveld, F. G. & Papantonis, A. Forces driving the three-dimensional folding of eukaryotic genomes. Mol. Syst. Biol. 14, e8214 (2018).

    PubMed  PubMed Central  Google Scholar 

  88. 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 

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

    CAS  PubMed  Google Scholar 

  90. Crane, E. et al. Condensin-driven remodelling of X chromosome topology during dosage compensation. Nature 523, 240–244 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  91. Dali, R. & Blanchette, M. A critical assessment of topologically associating domain prediction tools. Nucleic Acids Res. 45, 2994–3005 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  92. Zufferey, M., Tavernari, D., Oricchio, E. & Ciriello, G. Comparison of computational methods for the identification of topologically associating domains. Genome Biol. 19, 217 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  93. Forcato, M. et al. Comparison of computational methods for Hi-C data analysis. Nat. Methods 14, 679–685 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  94. Beagan, J. A. & Phillips-Cremins, J. E. On the existence and functionality of topologically associating domains. Nat. Genet. 52, 8–16 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  95. de Wit, E. TADs as the caller calls them. J. Mol. Biol. 432, 638–642 (2020).

    PubMed  Google Scholar 

  96. Dowen, J. M. et al. Control of cell identity genes occurs in insulated neighborhoods in mammalian chromosomes. Cell 159, 374–387 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  97. Gómez-Marín, C. et al. Evolutionary comparison reveals that diverging CTCF sites are signatures of ancestral topological associating domains borders. Proc. Natl Acad. Sci. USA 112, 7542 (2015).

    PubMed  PubMed Central  Google Scholar 

  98. Hanssen, L. L. P. et al. Tissue-specific CTCF–cohesin-mediated chromatin architecture delimits enhancer interactions and function in vivo. Nat. Cell Biol. 19, 952–961 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  99. Lupiáñez, D. G. et al. Disruptions of topological chromatin domains cause pathogenic rewiring of gene-enhancer interactions. Cell 161, 1012–1025 (2015).

    PubMed  PubMed Central  Google Scholar 

  100. Narendra, V. et al. CTCF establishes discrete functional chromatin domains at the Hox clusters during differentiation. Science 347, 1017–1021 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  101. Symmons, O. et al. The Shh topological domain facilitates the action of remote enhancers by reducing the effects of genomic distances. Dev. Cell 39, 529–543 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  102. Guo, Y. et al. CRISPR inversion of CTCF sites alters genome topology and enhancer/promoter function. Cell 162, 900–910 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  103. de Wit, E. et al. CTCF binding polarity determines chromatin looping. Mol. Cell 60, 676–684 (2015).

    PubMed  Google Scholar 

  104. Sanborn, A. L. et al. Chromatin extrusion explains key features of loop and domain formation in wild-type and engineered genomes. Proc. Natl Acad. Sci. USA 112, E6456–E6465 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. Paliou, C. et al. Preformed chromatin topology assists transcriptional robustness of Shh during limb development. Proc. Natl Acad. Sci. USA 116, 12390–12399 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  106. Fudenberg, G. et al. Formation of chromosomal domains by loop extrusion. Cell Rep. 15, 2038–2049 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  107. Hansen, A. S., Pustova, I., Cattoglio, C., Tjian, R. & Darzacq, X. CTCF and cohesin regulate chromatin loop stability with distinct dynamics. eLife 6, e25776 (2017).

    PubMed  PubMed Central  Google Scholar 

  108. Stevens, T. J. et al. 3D structures of individual mammalian genomes studied by single-cell Hi-C. Nature 544, 59–64 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  Google Scholar 

  110. Flyamer, I. M. et al. Single-nucleus Hi-C reveals unique chromatin reorganization at oocyte-to-zygote transition. Nature 544, 110–114 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  111. Gassler, J. et al. A mechanism of cohesin-dependent loop extrusion organizes zygotic genome architecture. EMBO J. 36, 3600–3618 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  112. Bintu, B. et al. Super-resolution chromatin tracing reveals domains and cooperative interactions in single cells. Science 362, eaau1783 (2018).

    PubMed  PubMed Central  Google Scholar 

  113. 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 

  114. Rubio, E. D. et al. CTCF physically links cohesin to chromatin. Proc. Natl Acad. Sci. USA 105, 8309 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  115. Stedman, W. et al. Cohesins localize with CTCF at the KSHV latency control region and at cellular c-myc and H19/Igf2 insulators. EMBO J. 27, 654–666 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  116. Fudenberg, G., Abdennur, N., Imakaev, M., Goloborodko, A. & Mirny, L. A. Emerging evidence of chromosome folding by loop extrusion. Cold Spring Harb. Symp. Quant. Biol. 82, 45–55 (2017).

    PubMed  Google Scholar 

  117. Rao, S. S. P. et al. Cohesin loss eliminates all loop domains. Cell 171, 305–320.e324 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  118. Wutz, G. et al. Topologically associating domains and chromatin loops depend on cohesin and are regulated by CTCF, WAPL, and PDS5 proteins. EMBO J. 36, 3573–3599 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  119. Schwarzer, W. et al. Two independent modes of chromatin organization revealed by cohesin removal. Nature 551, 51–56 (2017).

    PubMed  PubMed Central  Google Scholar 

  120. Haarhuis, J. H. I. et al. The cohesin release factor WAPL restricts chromatin loop extension. Cell 169, 693–707.e614 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  121. Li, Y. et al. The structural basis for cohesin–CTCF-anchored loops. Nature 578, 472–476 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  122. Davidson, I. F. et al. DNA loop extrusion by human cohesin. Science 366, 1338 (2019).

    CAS  PubMed  Google Scholar 

  123. Kim, Y., Shi, Z., Zhang, H., Finkelstein, I. J. & Yu, H. Human cohesin compacts DNA by loop extrusion. Science 366, 1345 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  124. Golfier, S., Quail, T., Kimura, H. & Brugués, J. Cohesin and condensin extrude DNA loops in a cell cycle-dependent manner. eLife 9, e53885 (2020).

    PubMed  PubMed Central  Google Scholar 

  125. Phillips-Cremins, J. E. et al. Architectural protein subclasses shape 3D organization of genomes during lineage commitment. Cell 153, 1281–1295 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  126. Abramo, K. et al. A chromosome folding intermediate at the condensin-to-cohesin transition during telophase. Nat. Cell Biol. 21, 1393–1402 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  127. Zhang, H. et al. Chromatin structure dynamics during the mitosis-to-G1 phase transition. Nature 576, 158–162 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  128. Nagano, T. et al. Cell-cycle dynamics of chromosomal organization at single-cell resolution. Nature 547, 61–67 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  129. Rhodes, J. D. P. et al. Cohesin disrupts polycomb-dependent chromosome interactions in embryonic stem cells. Cell Rep. 30, 820–835.e810 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  130. Nuebler, J., Fudenberg, G., Imakaev, M., Abdennur, N. & Mirny, L. A. Chromatin organization by an interplay of loop extrusion and compartmental segregation. Proc. Natl Acad. Sci. USA 115, E6697 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  131. Benabdallah, N. S. et al. Decreased enhancer-promoter proximity accompanying enhancer activation. Mol. Cell 76, 473–484.e477 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  132. Symmons, O. et al. Functional and topological characteristics of mammalian regulatory domains. Genome Res. 24, 390–400 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  133. Dekker, J. & Mirny, L. The 3D genome as moderator of chromosomal communication. Cell 164, 1110–1121 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  134. Chang, L.-H., Ghosh, S. & Noordermeer, D. TADs and their borders: free movement or building a wall? J. Mol. Biol. 432, 643–652 (2020).

    CAS  PubMed  Google Scholar 

  135. Vian, L. et al. The energetics and physiological impact of cohesin extrusion. Cell 173, 1165–1178.e1120 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  136. Barrington, C. et al. Enhancer accessibility and CTCF occupancy underlie asymmetric TAD architecture and cell type specific genome topology. Nat. Commun. 10, 2908 (2019).

    PubMed  PubMed Central  Google Scholar 

  137. Williamson, I. et al. Developmentally regulated Shh expression is robust to TAD perturbations. Development 146, dev179523 (2019).

    CAS  PubMed  Google Scholar 

  138. Franke, M. et al. Formation of new chromatin domains determines pathogenicity of genomic duplications. Nature 538, 265–269 (2016).

    CAS  PubMed  Google Scholar 

  139. Kragesteen, B. K. et al. Dynamic 3D chromatin architecture contributes to enhancer specificity and limb morphogenesis. Nat. Genet. 50, 1463–1473 (2018).

    CAS  PubMed  Google Scholar 

  140. Despang, A. et al. Functional dissection of the Sox9–Kcnj2 locus identifies nonessential and instructive roles of TAD architecture. Nat. Genet. 51, 1263–1271 (2019).

    CAS  PubMed  Google Scholar 

  141. Kraft, K. et al. Serial genomic inversions induce tissue-specific architectural stripes, gene misexpression and congenital malformations. Nat. Cell Biol. 21, 305–310 (2019).

    CAS  PubMed  Google Scholar 

  142. Oudelaar, A. M. et al. Dynamics of the 4D genome during in vivo lineage specification and differentiation. Nat. Commun. 11, 2722 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  143. Cullen, K. E., Kladde, M. P. & Seyfred, M. A. Interaction between transcription regulatory regions of prolactin chromatin. Science 261, 203 (1993).

    CAS  PubMed  Google Scholar 

  144. Ptashne, M. Gene regulation by proteins acting nearby and at a distance. Nature 322, 697–701 (1986).

    CAS  PubMed  Google Scholar 

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

    CAS  PubMed  Google Scholar 

  146. Carter, D., Chakalova, L., Osborne, C. S., Dai, Y.-F. & Fraser, P. Long-range chromatin regulatory interactions in vivo. Nat. Genet. 32, 623–626 (2002).

    CAS  PubMed  Google Scholar 

  147. Vakoc, C. R. et al. Proximity among distant regulatory elements at the β-globin locus requires GATA-1 and FOG-1. Mol. Cell 17, 453–462 (2005).

    CAS  PubMed  Google Scholar 

  148. Drissen, R. et al. The active spatial organization of the β-globin locus requires the transcription factor EKLF. Genes Dev. 18, 2485–2490 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  149. 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 

  150. Song, S.-H., Hou, C. & Dean, A. A positive role for NLI/Ldb1 in long-range β-globin locus control region function. Mol. Cell 28, 810–822 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  151. Deng, W. et al. Reactivation of developmentally silenced globin genes by forced chromatin looping. Cell 158, 849–860 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  152. Krivega, I., Dale, R. K. & Dean, A. Role of LDB1 in the transition from chromatin looping to transcription activation. Genes Dev. 28, 1278–1290 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  153. Kim, S. & Shendure, J. Mechanisms of interplay between transcription factors and the 3D genome. Mol. Cell 76, 306–319 (2019).

    CAS  PubMed  Google Scholar 

  154. Weintraub, A. S. et al. YY1 is a structural regulator of enhancer-promoter loops. Cell 171, 1573–1588.e1528 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  155. Beagan, J. A. et al. YY1 and CTCF orchestrate a 3D chromatin looping switch during early neural lineage commitment. Genome Res. 27, 1139–1152 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  156. Kagey, M. H. et al. Mediator and cohesin connect gene expression and chromatin architecture. Nature 467, 430–435 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  157. El Khattabi, L. et al. A pliable mediator acts as a functional rather than an architectural bridge between promoters and enhancers. Cell 178, 1145–1158.e1120 (2019).

    CAS  PubMed  Google Scholar 

  158. Jaeger, M. G. et al. Selective mediator dependence of cell-type-specifying transcription. Nat. Genet. 52, 719–727 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

  159. Morgan, S. L. et al. Manipulation of nuclear architecture through CRISPR-mediated chromosomal looping. Nat. Commun. 8, 15993 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  160. Kim, J. H. et al. LADL: light-activated dynamic looping for endogenous gene expression control. Nat. Methods 16, 633–639 (2019).

    PubMed  PubMed Central  Google Scholar 

  161. Haberle, V. et al. Transcriptional cofactors display specificity for distinct types of core promoters. Nature 570, 122–126 (2019).

    CAS  PubMed  Google Scholar 

  162. Schmidt, D. et al. A CTCF-independent role for cohesin in tissue-specific transcription. Genome Res. 20, 578–588 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  163. Cuartero, S. et al. Control of inducible gene expression links cohesin to hematopoietic progenitor self-renewal and differentiation. Nat. Immunol. 19, 932–941 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  164. Chiariello, A. M. et al. A dynamic folded hairpin conformation is associated with α-globin activation in erythroid cells. Cell Rep. 30, 2125–2135.e2125 (2020).

    CAS  PubMed  Google Scholar 

  165. Buckle, A., Brackley, C. A., Boyle, S., Marenduzzo, D. & Gilbert, N. Polymer simulations of heteromorphic chromatin predict the 3D folding of complex genomic loci. Mol. Cell 72, 786–797.e711 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  166. Zheng, H. & Xie, W. The role of 3D genome organization in development and cell differentiation. Nat. Rev. Mol. Cell Biol. 34, 903–916 (2019).

    Google Scholar 

  167. Dixon, J. R. et al. Chromatin architecture reorganization during stem cell differentiation. Nature 518, 331–336 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  168. Bonev, B. et al. Multiscale 3D genome rewiring during mouse neural development. Cell 171, 557–572.e524 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  169. Stadhouders, R. et al. Transcription factors orchestrate dynamic interplay between genome topology and gene regulation during cell reprogramming. Nat. Genet. 50, 238–249 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  170. Brown, J. M. et al. A tissue-specific self-interacting chromatin domain forms independently of enhancer-promoter interactions. Nat. Commun. 9, 376 (2018).

    Google Scholar 

  171. Jin, F. et al. A high-resolution map of the three-dimensional chromatin interactome in human cells. Nature 503, 290–294 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  172. Ghavi-Helm, Y. et al. Enhancer loops appear stable during development and are associated with paused polymerase. Nature 513, 89–100 (2014).

    Google Scholar 

  173. Andrey, G. et al. A switch between topological domains underlies HoxD genes collinearity in mouse limbs. Science 340, 1234167 (2013).

    PubMed  Google Scholar 

  174. Rubin, A. J. et al. Lineage-specific dynamic and pre-established enhancer–promoter contacts cooperate in terminal differentiation. Nat. Genet. 49, 1522–1528 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  175. de Laat, W. & Duboule, D. Topology of mammalian developmental enhancers and their regulatory landscapes. Nature 502, 499–506 (2013).

    PubMed  Google Scholar 

  176. Freire-Pritchett, P. et al. Global reorganisation of cis-regulatory units upon lineage commitment of human embryonic stem cells. eLife 6, e21926 (2017).

    PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  Google Scholar 

  178. Vernimmen, D., Gobbi, M. D., 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 

  179. Williamson, I., Lettice, L. A., Hill, R. E. & Bickmore, W. A. Shh and ZRS enhancer colocalisation is specific to the zone of polarising activity. Development 143, 2994–3001 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  180. Boettiger, A. N. et al. Super-resolution imaging reveals distinct chromatin folding for different epigenetic states. Nature 529, 418–422 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  181. Mateo, L. J. et al. Visualizing DNA folding and RNA in embryos at single-cell resolution. Nature 568, 49–54 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  182. Wang, S. et al. Spatial organization of chromatin domains and compartments in single chromosomes. Science 353, 598 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  183. Cardozo Gizzi, A. M. et al. Microscopy-based chromosome conformation capture enables simultaneous visualization of genome organization and transcription in intact organisms. Mol. Cell 74, 212–222.e215 (2019).

    CAS  PubMed  Google Scholar 

  184. Szabo, Q. et al. TADs are 3D structural units of higher-order chromosome organization in Drosophila. Sci. Adv. 4, eaar8082 (2018).

    PubMed  PubMed Central  Google Scholar 

  185. Fabre, P. J. et al. Large scale genomic reorganization of topological domains at the HoxD locus. Genome Biol. 18, 149 (2017).

    PubMed  PubMed Central  Google Scholar 

  186. Banani, S. F., Lee, H. O., Hyman, A. A. & Rosen, M. K. Biomolecular condensates: organizers of cellular biochemistry. Nat. Rev. Mol. Cell Biol. 18, 285–298 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  187. Alberti, S. Phase separation in biology. Curr. Biol. 27, R1097–R1102 (2017).

    CAS  PubMed  Google Scholar 

  188. Boeynaems, S. et al. Protein phase separation: a new phase in cell biology. Trends Cell Biol. 28, 420–435 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  189. Sabari, B. R., Dall’Agnese, A. & Young, R. A. Biomolecular condensates in the nucleus. Trends Biochem. Sci. https://doi.org/10.1016/j.tibs.2020.06.007 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  Google Scholar 

  191. Beagrie, R. A. et al. Complex multi-enhancer contacts captured by genome architecture mapping. Nature 543, 519–524 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  192. Cho, W.-K. et al. Mediator and RNA polymerase II clusters associate in transcription-dependent condensates. Science 361, 412 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  193. Cisse, I. I. et al. Real-time dynamics of RNA polymerase II clustering in live human cells. Science 341, 664 (2013).

    CAS  PubMed  Google Scholar 

  194. Jackson, D. A., Hassan, A. B., Errington, R. J. & Cook, P. R. Visualization of focal sites of transcription within human nuclei. EMBO J. 12, 1059–1065 (1993).

    CAS  PubMed  PubMed Central  Google Scholar 

  195. Brown, J. M. et al. Association between active genes occurs at nuclear speckles and is modulated by chromatin environment. J. Cell Biol. 182, 1083–1097 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  196. Chen, Y. et al. Mapping 3D genome organization relative to nuclear compartments using TSA-Seq as a cytological ruler. J. Cell Biol. 217, 4025–4048 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  197. Quinodoz, S. A. et al. Higher-order inter-chromosomal hubs shape 3D genome organization in the nucleus. Cell 174, 744–757.e724 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  198. 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 

  199. Denholtz, M. et al. Long-range chromatin contacts in embryonic stem cells reveal a role for pluripotency factors and polycomb proteins in genome organization. Cell Stem Cell 13, 602–616 (2013).

    CAS  PubMed  Google Scholar 

  200. Fudenberg, G., Kelley, D. R. & Pollard, K. S. Predicting 3D genome folding from DNA sequence with Akita. Nat. Methods https://doi.org/10.1038/s41592-020-0958-x (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  201. Schwessinger, R. et al. DeepC: predicting 3D genome folding using megabase-scale transfer learning. Nat. Methods https://doi.org/10.1038/s41592-020-0960-3 (2020).

  202. Zhao, Z. et al. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat. Genet. 38, 1341–1347 (2006).

    CAS  PubMed  Google Scholar 

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

    CAS  PubMed  Google Scholar 

  204. Davies, J. O. J. et al. Multiplexed analysis of chromosome conformation at vastly improved sensitivity. Nat. Methods 86, 1202–1210 (2015).

    Google Scholar 

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

    CAS  PubMed  Google Scholar 

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

    CAS  PubMed  Google Scholar 

  207. Übelmesser, N. & Papantonis, A. Technologies to study spatial genome organization: beyond 3C. Brief. Funct. Genomics 18, 395–401 (2019).

    PubMed  Google Scholar 

  208. Brant, L. et al. Exploiting native forces to capture chromosome conformation in mammalian cell nuclei. Mol. Syst. Biol. 12, 891 (2016).

    PubMed  PubMed Central  Google Scholar 

  209. Redolfi, J. et al. DamC reveals principles of chromatin folding in vivo without crosslinking and ligation. Nat. Struct. Mol. Biol. 26, 471–480 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  210. Finn, E. H. et al. Extensive heterogeneity and intrinsic variation in spatial genome organization. Cell 176, 1502–1515.e1510 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  211. Nguyen, H. Q. et al. 3D mapping and accelerated super-resolution imaging of the human genome using in situ sequencing. Nat. Methods 17, 822–832 (2020).

    CAS  PubMed  PubMed Central  Google Scholar 

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

    CAS  PubMed  Google Scholar 

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Acknowledgements

The authors thank R. A. Beagrie, J. M. Brown and M. T. Kassouf for insightful comments on the manuscript. The authors apologize to colleagues whose important studies they were unable to cite due to space constraints. This work was supported by the Max Planck Society (A.M.O.) and the UK Medical Research Council (D.H.; MR/T014067/1).

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Glossary

Epigenetic

Pertains to changes in chromatin that register, signal or perpetuate altered activity states without changing the primary DNA sequence.

Chromatin

The complex of DNA and proteins that makes up chromosomes; chromatin consists of nucleosomes formed of ~150 bp of DNA wrapped around a histone octamer, which can be further packaged into higher-order structures.

Euchromatin

A relatively loosely packaged form of chromatin that is enriched in genes that are actively transcribed or poised for transcription.

Facultative heterochromatin

Regions of chromatin that are densely packaged and transcriptionally silent but may lose their condensed state and become transcriptionally active.

Constitutive heterochromatin

Regions of permanently densely packaged and transcriptionally silent chromatin that are found at specific, highly repetitive regions of the genome, such as telomeres and centromeres.

Pre-initiation complex

A large complex of proteins, including RNA polymerase II and its associated general transcription factors, which is necessary for the transcription of protein-coding genes.

TATA box

Named after its conserved DNA sequence, the TATA box is a non-coding DNA sequence found in many eukaryotic core promoters that recruits the pre-initiation complex to initiate transcription.

CpG islands

Regions of the genome (~300–3,000 bp) that contain a large number of CpG dinucleotides and are associated with ~40–70% of mammalian gene promoters.

Mediator complex

A large protein complex that acts as a key transcriptional co-activator by communicating signals from transcription factors to RNA polymerase II to control its activity.

Nuclear speckles

Non-membrane-bound subdomains located in the interchromatin regions of the nucleus of mammalian cells that are enriched in splicing factors and other mRNA-processing proteins.

Phase separation

The process by which substances in a mixture become separated in two distinct phases, as occurs in a mixture of oil and water.

Polycomb

The Polycomb system involves various protein complexes, including Polycomb repressive complex 1 (PRC1) and PRC2, which act as transcriptional repressors with a key role in epigenetic silencing during differentiation and development.

Polycomb bodies

Foci of Polycomb group proteins in the nucleus that are involved in both genome organization and repression of gene expression.

Molecular condensates

Non-membrane-bound subcompartments in the nucleoplasm or cytoplasm that are strongly enriched in or depleted of specific proteins or nucleic acids.

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Oudelaar, A.M., Higgs, D.R. The relationship between genome structure and function. Nat Rev Genet 22, 154–168 (2021). https://doi.org/10.1038/s41576-020-00303-x

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