Studies of 3D chromatin organization have suggested that chromosomes are hierarchically organized into large compartments composed of smaller domains called topologically associating domains (TADs). Recent evidence suggests that compartments are smaller than previously thought and that the transcriptional or chromatin state is responsible for interactions leading to the formation of small compartmental domains in all organisms. In vertebrates, CTCF forms loop domains, probably via an extrusion process involving cohesin. CTCF loops cooperate with compartmental domains to establish the 3D organization of the genome. The continuous extrusion of the chromatin fibre by cohesin may also be responsible for the establishment of enhancer–promoter interactions and stochastic aspects of the transcription process. These observations suggest that the 3D organization of the genome is an emergent property of chromatin and its components, and thus may not be only a determinant but also a consequence of its function.
Subscribe to Journal
Get full journal access for 1 year
only $21.58 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Fraser, J., Williamson, I., Bickmore, W. A. & Dostie, J. An overview of genome organization and how we got there: from FISH to Hi-C. Microbiol. Mol. Biol. Rev. 79, 347–372 (2015).
Stevens, T. J. et al. 3D structures of individual mammalian genomes studied by single-cell Hi-C. Nature 544, 59–64 (2017).
Nagano, T. et al. Single-cell Hi-C reveals cell-to-cell variability in chromosome structure. Nature 502, 59–64 (2013).
Beliveau, B. J. et al. Single-molecule super-resolution imaging of chromosomes and in situ haplotype visualization using Oligopaint FISH probes. Nat. Commun. 6, 7147 (2015).
Ni, Y. et al. Super-resolution imaging of a 2.5 kb non-repetitive DNA in situ in the nuclear genome using molecular beacon probes. eLife 6, e21660 (2017).
Nagano, T. et al. Cell-cycle dynamics of chromosomal organization at single-cell resolution. Nature 547, 61–67 (2017).
Pegoraro, G. & Misteli, T. High-throughput imaging for the discovery of cellular mechanisms of disease. Trends Genet. 33, 604–615 (2017).
Lieberman-Aiden, E. et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009).
Lajoie, B. R., Dekker, J. & Kaplan, N. The hitchhiker’s guide to Hi-C analysis: practical guidelines. Methods 72, 65–75 (2015).
Dixon, J. R. et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485, 376–380 (2012).
Sexton, T. et al. Three-dimensional folding and functional organization principles of the Drosophila genome. Cell 148, 458–472 (2012).
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).
Nora, E. P. et al. Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 485, 381–385 (2012).
Bonev, B. & Cavalli, G. Organization and function of the 3D genome. Nat. Rev. Genet. 17, 661–678 (2016).
Schmitt, A. D., Hu, M. & Ren, B. Genome-wide mapping and analysis of chromosome architecture. Nat. Rev. Mol. Cell. Biol. 17, 743–755 (2016).
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).
Rowley, M. J. et al. Evolutionarily Conserved Principles Predict 3D Chromatin Organization. Mol. Cell 67, 837–852 (2017).
Rao, S. et al. Cohesin loss eliminates all loop domains, leading to links among superenhancers and downregulation of nearby genes. Preprint at BioRxiv https://doi.org/10.1101/139782 (2017).
Dong, P. et al. 3D chromatin architecture of large plant genomes determined bylocal A/B compartments. Mol. Plant 10, 1497–1509 (2017).
Haddad, N., Jost, D. & Vaillant, C. Perspectives: using polymer modeling to understand the formation and function of nuclear compartments. Chromosome Res. 25, 35–50 (2017).
Huang, J., Marco, E., Pinello, L. & Yuan, G.-C. Predicting chromatin organization using histone marks. Genome Biol. 16, 162 (2015).
Di Pierro, M., Cheng, R. R., Lieberman Aiden, E., Wolynes, P. G. & Onuchic, J. N. De novo prediction of human chromosome structures: epigenetic marking patterns encode genome architecture. Proc. Natl Acad. Sci. USA 114, 12126–12131 (2017).
Wang, X., Brandão, H. B., Le, T. B. K., Laub, M. T. & Rudner, D. Z. Bacillus subtilis SMC complexes juxtapose chromosome arms as they travel from origin to terminus. Science 355, 524–527 (2017).
Le, T. B. K., Imakaev, M. V., Mirny, L. A. & Laub, M. T. High-resolution mapping of the spatial organization of a bacterial chromosome. Science 342, 731–734 (2013).
Li, L. et al. Widespread rearrangement of 3D chromatin organization underlies polycomb-mediated stress-induced silencing. Mol. Cell 58, 216–231 (2015).
Hug, C. B., Grimaldi, A. G., Kruse, K. & Vaquerizas, J. M. Chromatin Architecture Emerges during Zygotic Genome Activation Independent of Transcription. Cell 169, 216–228 (2017).
Bensaude, O. Inhibiting eukaryotic transcription. Which compound to choose? How to evaluate its activity?: Which compound to choose? How to evaluate its activity? Transcription 2, 103–108 (2011).
Kaaij, L. J. T., van der Weide, R. H., Ketting, R. F. & de Wit, E. Systemic loss and gain of chromatin architecture throughout zebrafish development. Cell Rep. 24, 1–10 (2018).
Du, Z. et al. Allelic reprogramming of 3D chromatin architecture during early mammalian development. Nature 547, 232–235 (2017).
Ke, Y. et al. 3D chromatin structures of mature gametes and structural reprogramming during mammalian embryogenesis. Cell 170, 367–381 (2017).
Naumova, N. et al. Organization of the mitotic chromosome. Science 342, 948–953 (2013).
Jung, Y. H. et al. Chromatin states in mouse sperm correlate with embryonic and adult regulatory landscapes. Cell Rep. 18, 1366–1382 (2017).
Battulin, N. et al. Comparison of the three-dimensional organization of sperm and fibroblast genomes using the Hi-C approach. Genome Biol. 16, 77 (2015).
Cubeñas-Potts, C. & Corces, V. G. Architectural proteins, transcription, and the three-dimensional organization of the genome. FEBS Lett. 589, 2923–2930 (2015).
Cubeñas-Potts, C. et al. Different enhancer classes in Drosophila bind distinct architectural proteins and mediate unique chromatin interactions and 3D architecture. Nucleic Acids Res. 45, 1714–1730 (2016).
Harlen, K. M. & Churchman, L. S. The code and beyond: transcription regulation by the RNA polymerase II carboxy-terminal domain. Nat. Rev. Mol. Cell. Biol. 18, 263–273 (2017).
Hnisz, D., Shrinivas, K., Young, R. A., Chakraborty, A. K. & Sharp, P. A. A. Phase separation model for transcriptional control. Cell 169, 13–23 (2017).
Larson, A. G. et al. Liquid droplet formation by HP1α suggests a role for phase separation in heterochromatin. Nature 547, 236–240 (2017).
Lin, Y., Currie, S. L. & Rosen, M. K. Intrinsically disordered sequences enable modulation of protein phase separation through distributed tyrosine motifs. J. Biol. Chem. 292, 19110–19120 (2017).
van der Lee, R. et al. Classification of intrinsically disordered regions and proteins. Chem. Rev. 114, 6589–6631 (2014).
Lu, H. et al. Phase-separation mechanism for C-terminal hyperphosphorylation of RNA polymerase II. Nature 558, 318–323 (2018).
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, 3503–3512 (2016).
Tang, Z. et al. CTCF-mediated human 3D genome architecture reveals chromatin topology for transcription. Cell 163, 1611–1627 (2015).
Guo, Y. et al. CRISPR inversion of CTCF sites alters genome topology and enhancer/promoter function. Cell 162, 900–910 (2015).
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).
Fudenberg, G. et al. Formation of chromosomal domains by loop extrusion. Cell Rep. 15, 2038–2049 (2016).
Nichols, M. H. & Corces, V. G. A. CTCF code for 3D genome architecture. Cell 162, 703–705 (2015).
Nasmyth, K. Disseminating the genome: joining, resolving, and separating sister chromatids during mitosis and meiosis. Annu. Rev. Genet. 35, 673–745 (2001).
Alipour, E. & Marko, J. F. Self-organization of domain structures by DNA-loop-extruding enzymes. Nucleic Acids Res. 40, 11202–11212 (2012).
Stigler, J., Çamdere, G. Ö., Koshland, D. E. & Greene, E. C. Single-molecule imaging reveals a collapsed conformational state for DNA-bound cohesin. Cell Rep. 15, 988–998 (2016).
Davidson, I. F. et al. Rapid movement and transcriptional re-localization of human cohesin on DNA. EMBO J. 35, 2671–2685 (2016).
Kanke, M., Tahara, E., Huis In’t Veld, P. J. & Nishiyama, T. Cohesin acetylation and Wapl-Pds5 oppositely regulate translocation of cohesin along DNA. EMBO J. 35, 2686–2698 (2016).
Ganji, M. et al. Real-time imaging of DNA loop extrusion by condensin. Science 360, 102–105 (2018).
Terakawa, T. et al. The condensin complex is a mechanochemical motor that translocates along DNA. Science 358, 672–676 (2017).
Murayama, Y., Samora, C. P., Kurokawa, Y., Iwasaki, H. & Uhlmann, F. Establishment of DNA-DNA interactions by the cohesin ring. Cell 172, 465–477 (2018).
Nagy, G. et al. Motif oriented high-resolution analysis of ChIP-seq data reveals the topological order of CTCF and cohesin proteins on DNA. BMC Genomics 17, 637 (2016).
Schwarzer, W. et al. Two independent modes of chromatin organization revealed by cohesin removal. Nature 551, 51–56 (2017).
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).
Haarhuis, J. H. I. et al. The cohesin release factor WAPL restricts chromatin loop extension. Cell 169, 693–707 (2017).
Gassler, J. et al. A mechanism of cohesin-dependent loop extrusion organizes zygotic genome architecture. EMBO J. 36, 3600–3618 (2017).
Fu, Y., Sinha, M., Peterson, C. L. & Weng, Z. The insulator binding protein CTCF positions 20 nucleosomes around its binding sites across the human genome. PLOS Genet. 4, e1000138 (2008).
Mawhinney, M. T. et al. CTCF-induced circular DNA complexes observed by atomic force microscopy. J. Mol. Biol. 430, 759–776 (2018).
Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944 (2017).
Kubo, N. et al. Preservation of chromatin organization after acute loss of CTCF in mouse embryonic stem cells. Preprint at BioRxiv. https://doi.org/10.1101/118737 (2017).
Phanstiel, D. H. et al. Static and dynamic DNA loops form AP-1-bound activation hubs during macrophage development. Mol. Cell 67, 1037–1048 (2017).
Bonev, B. et al. Multiscale 3D genome rewiring during mouse neural development. Cell 171, 557–572 (2017).
Gibcus, J. H. et al. A pathway for mitotic chromosome formation. Science 359, eaao6135 (2018).
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).
Tran, N. T., Laub, M. T. & Le, T. B. K. SMC progressively aligns chromosomal arms in Caulobacter crescentus but is antagonized by convergent transcription. Cell Rep. 20, 2057–2071 (2017).
Brackley, C. A. et al. Nonequilibrium chromosome looping via molecular slip links. Phys. Rev. Lett. 119, 138101 (2017).
Flamholz, A., Phillips, R. & Milo, R. The quantified cell. Mol. Biol. Cell 25, 3497–3500 (2014).
Vian, L. et al. The energetics and physiological impact of cohesin extrusion. Cell 173, 1165–1178 (2018).
Ocampo-Hafalla, M., Muñoz, S., Samora, C. P. & Uhlmann, F. Evidence for cohesin sliding along budding yeast chromosomes. Open Biol. 6, 150178 (2016).
Bausch, C. et al. Transcription alters chromosomal locations of cohesin in Saccharomyces cerevisiae. Mol. Cell. Biol. 27, 8522–8532 (2007).
Busslinger, G. A. et al. Cohesin is positioned in mammalian genomes by transcription, CTCF and Wapl. Nature 544, 503–507 (2017).
Racko, D., Benedetti, F., Dorier, J. & Stasiak, A. Transcription-induced supercoiling as the driving force of chromatin loop extrusion during formation of TADs in interphase chromosomes. Nucleic Acids Res. 46, 1648–1660 (2017).
Jonkers, I. & Lis, J. T. Getting up to speed with transcription elongation by RNA polymerase II. Nat. Rev. Mol. Cell. Biol. 16, 167–177 (2015).
Moore, J. M. et al. Loss of maternal CTCF is associated with peri-implantation lethality of Ctcf null embryos. PLOS ONE 7, e34915 (2012).
Dowen, J. M. et al. Control of cell identity genes occurs in insulated neighborhoods in mammalian chromosomes. Cell 159, 374–387 (2014).
Hnisz, D., Day, D. S. & Young, R. A. Insulated neighborhoods: structural and functional units of mammalian gene control. Cell 167, 1188–1200 (2016).
Wan, L.-B. et al. Maternal depletion of CTCF reveals multiple functions during oocyte and preimplantation embryo development. Development 135, 2729–2738 (2008).
Gregor, A. et al. De novo mutations in the genome organizer CTCF cause intellectual disability. Am. J. Hum. Genet. 93, 124–131 (2013).
Kemp, C. J. et al. CTCF haploinsufficiency destabilizes DNA methylation and predisposes to cancer. Cell Rep. 7, 1020–1029 (2014).
Katainen, R. et al. CTCF/cohesin-binding sites are frequently mutated in cancer. Nat. Genet. 47, 818–821 (2015).
Narendra, V. et al. CTCF establishes discrete functional chromatin domains at the Hox clusters during differentiation. Science 347, 1017–1021 (2015).
de Wit, E. et al. CTCF binding polarity determines chromatin looping. Mol. Cell 60, 676–684 (2015).
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).
Kojic, A. et al. Distinct roles of cohesin-SA1 and cohesin-SA2 in 3D chromosome organization. Nat. Struct. Mol. Biol. 25, 496–504 (2018).
Fukaya, T., Lim, B. & Levine, M. Enhancer control of transcriptional bursting. Cell 166, 358–368 (2016).
Weintraub, A. S. et al. YY1 is a structural regulator of enhancer-promoter loops. Cell 171, 1573–1588 (2017).
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).
Pan, X. et al. YY1 controls Igκ repertoire and B cell development, and localizes with condensin on the Igκ locus. EMBO J. 32, 1168–1182 (2013).
Flyamer, I. M. et al. Single-nucleus Hi-C reveals unique chromatin reorganization at oocyte-to-zygote transition. Nature 544, 110–114 (2017).
Gligoris, T. & Löwe, J. Structural insights into ring formation of cohesin and related Smc complexes. Trends Cell Biol. 26, 680–693 (2016).
Work in the authors’ laboratory is supported by US Public Health Service Award R01 GM035463 (V.G.C.) and Pathway to Independence Award K99/R00 GM127671 (M.J.R.) from the US National Institutes of Health (NIH). The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH.
The authors declare no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Chromatin interaction analysis by paired-end tag sequencing (ChIA-PET) utilizes chromatin immunoprecipitation followed by proximity ligation to identify chromatin interactions between loci bound by a protein of interest.
- ChIP–exo and ChIP–nexus
Chromatin immunoprecipitation (ChIP) followed by exonuclease digestion (ChIP–exo) is a technique that is used in place of standard chromatin immunoprecipitation followed by sequencing (ChIP–seq) to identify protein binding sites at higher resolution. The higher resolution is achieved because exonuclease treatment trims stretches of flanking DNA that are not directly bound by the protein of interest. ChIP–nexus utilizes a different library preparation strategy to reportedly improve signal compared with ChIP–exo.
- Compartmental domains
Domains in Hi-C data that are not formed by a CTCF loop and are formed instead by the segregation of active and inactive chromatin.
- CTCF loops
Point-to-point interactions between loci that coincide with CTCF and cohesin occupancy and often contain CTCF motifs in convergent orientation. These appear as bright punctae corresponding to high-frequency interactions in Hi-C contact maps.
- Directionality index
A common method of computationally identifying topologically associating domain (TAD) borders. A directionality is calculated for each binned genomic locus to describe the preference of interaction signal with bins on the right (positive directionality) or with bins on the left (negative directionality). TAD borders are defined at transitions between negative and positive directionality.
- Gene loop
A loop formed by interactions between the transcription start site and the transcription termination site.
- Global run-on sequencing
(GRO-seq). A method involving isolation of nascent transcripts and high-throughput sequencing to study active transcription genome-wide.
A method using proximity ligation and high-throughput sequencing to identify all interactions taking place throughout the genome.
- Loop extrusion
A model in which chromatin is pulled through the cohesin or condensin ring to form loops.
A method of labelling DNA using short fluorescently labelled oligonucleotides for high-resolution imaging of chromatin.
- Ordinary domains
Domains observed in Hi-C data that are not spanned by a CTCF loop. They are probably the same as compartmental domains.
(Stochastic optical reconstruction microscopy). Super-resolution imaging using individual photo-switchable fluorophores.
- Transcriptional states
The state of a locus based on the presence of chromatin-bound proteins or covalent histone modifications that correlate with gene silencing or active transcription.
- Transcription factory
A distinct nuclear location where RNA polymerase II (RNAPII) accumulates on the basis of the observation that components of the transcription complex can be detected as discrete foci by microscopy. The transcription factory hypothesis suggests that genes are recruited to these nuclear locations in order to be transcribed.
About this article
Cite this article
Rowley, M.J., Corces, V.G. Organizational principles of 3D genome architecture. Nat Rev Genet 19, 789–800 (2018). https://doi.org/10.1038/s41576-018-0060-8
Frontiers in Genetics (2020)
Estrogen receptor α (ERα)-binding super-enhancers drive key mediators that control uterine estrogen responses in mice
Journal of Biological Chemistry (2020)
Epigenetics & Chromatin (2020)
Science China Life Sciences (2020)