Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Review Article
  • Published:

New insights into genome folding by loop extrusion from inducible degron technologies

Nature Reviews Genetics volume 24, pages 73–85 (2023)Cite this article

Subjects

Abstract

Chromatin folds into dynamic loops that often span hundreds of kilobases and physically wire distant loci together for gene regulation. These loops are continuously created, extended and positioned by structural maintenance of chromosomes (SMC) protein complexes, such as condensin and cohesin, and their regulators, including CTCF, in a highly dynamic process known as loop extrusion. Genetic loss of extrusion factors is lethal, complicating their study. Inducible protein degradation technologies enable the depletion of loop extrusion factors within hours, leading to the rapid reconfiguration of chromatin folding. Here, we review how these technologies have changed our understanding of genome organization, upsetting long-held beliefs on its role in transcription. Finally, we examine recent models that attempt to reconcile observations after chronic versus acute perturbations, and discuss future developments in this rapidly developing field of research.

This is a preview of subscription content, access via your institution

Access options

Buy this article

  • Purchase on Springer Link
  • Instant access to full article PDF
Buy now

Prices may be subject to local taxes which are calculated during checkout

Fig. 1: Basic features of chromosome organization.
Fig. 2: Cohesin-mediated loop extrusion.
Fig. 3: Inducible degron technologies.
Fig. 4: Experimental strategies with inducible degron systems.
Fig. 5: Time-delayed effects of promoter–enhancer communication.

Similar content being viewed by others

References

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

    Article  CAS  Google Scholar 

  2. Hsieh, T.-H. S. et al. Mapping nucleosome resolution chromosome folding in yeast by Micro-C. Cell 162, 108–119 (2015).

    Article  CAS  Google Scholar 

  3. Bintu, B. et al. Super-resolution chromatin tracing reveals domains and cooperative interactions in single cells. Science 362, eaau1783 (2018). In this work, microscopy and DNA fluorescence in situ hybridization experiments were used to show that TAD structures are heterogeneous at the single-cell (and chromosome) level, and that cohesin degradation randomizes chromosome folding.

    Article  Google Scholar 

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

    Article  CAS  Google Scholar 

  5. Misteli, T. The self-organizing genome: principles of genome architecture and function. Cell 183, 28–45 (2020).

    Article  CAS  Google Scholar 

  6. Razin, S. V. & Ulianov, S. V. Gene functioning and storage within a folded genome. Cell. Mol. Biol. Lett. 22, 18 (2017).

    Article  Google Scholar 

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

    Article  CAS  Google Scholar 

  8. Gibcus, J. H. et al. A pathway for mitotic chromosome formation. Science 359, eaao6135 (2018). This work charts 3D genome structures leading up to pro-metaphase at high temporal resolution and after degradation of condensins resulting in a model for the formation of mitotic chromosomes.

    Article  Google Scholar 

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

    Article  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

  11. Solovei, I., Thanisch, K. & Feodorova, Y. How to rule the nucleus: divide et impera. Curr. Opin. Cell Biol. 40, 47–59 (2016).

    Article  CAS  Google Scholar 

  12. Bhat, P., Honson, D. & Guttman, M. Nuclear compartmentalization as a mechanism of quantitative control of gene expression. Nat. Rev. Mol. Cell Biol. 22, 653–670 (2021).

    Article  CAS  Google Scholar 

  13. Ferrai, C., Castro, I. J., de, Lavitas, L., Chotalia, M. & Pombo, A. Gene positioning. Cold Spring Harb. Perspect. Biol. 2, a000588 (2010).

    Article  Google Scholar 

  14. Hoskins, V. E., Smith, K. & Reddy, K. L. The shifting shape of genomes: dynamics of heterochromatin interactions at the nuclear lamina. Curr. Opin. Genet. Dev. 67, 163–173 (2021).

    Article  CAS  Google Scholar 

  15. van Steensel, B. & Belmont, A. S. Lamina-associated domains: links with chromosome architecture, heterochromatin, and gene repression. Cell 169, 780–791 (2017).

    Article  Google Scholar 

  16. Davidson, I. F. & Peters, J.-M. Genome folding through loop extrusion by SMC complexes. Nat. Rev. Mol. Cell Biol. 22, 445–464 (2021).

    Article  CAS  Google Scholar 

  17. Kim, Y. & Yu, H. Shaping of the 3D genome by the ATPase machine cohesin. Exp. Mol. Med. 52, 1891–1897 (2020).

    Article  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

  19. van Ruiten, M. S. & Rowland, B. D. On the choreography of genome folding: a grand pas de deux of cohesin and CTCF. Curr. Opin. Cell Biol. 70, 84–90 (2021).

    Article  Google Scholar 

  20. 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).

    Article  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

  22. Gomes, N. P. & Espinosa, J. M. Gene-specific repression of the p53 target gene PUMA via intragenic CTCF–cohesin binding. Genes Dev. 24, 1022–1034 (2010).

    Article  CAS  Google Scholar 

  23. Fudenberg, G. & Nora, E. P. Embryogenesis without CTCF in flies and vertebrates. Nat. Struct. Mol. Biol. 28, 774–776 (2021).

    Article  CAS  Google Scholar 

  24. Singh, V. P. & Gerton, J. L. Cohesin and human disease: lessons from mouse models. Curr. Opin. Cell Biol. 37, 9–17 (2015).

    Article  CAS  Google Scholar 

  25. Michaelis, C., Ciosk, R. & Nasmyth, K. Cohesins: chromosomal proteins that prevent premature separation of sister chromatids. Cell 91, 35–45 (1997).

    Article  CAS  Google Scholar 

  26. Busslinger, G. A. et al. Cohesin is positioned in mammalian genomes by transcription, CTCF and Wapl. Nature 544, 503–507 (2017).

    Article  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

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

    Article  Google Scholar 

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

    Article  CAS  Google Scholar 

  30. Calderon, L. et al. Cohesin-dependence of neuronal gene expression relates to chromatin loop length. eLife 11, e76539 (2022).

    Article  CAS  Google Scholar 

  31. Yatskevich, S., Rhodes, J. & Nasmyth, K. Organization of chromosomal DNA by SMC complexes. Annu. Rev. Genet. 53, 445–482 (2019).

    Article  CAS  Google Scholar 

  32. Nasmyth, K. Disseminating the genome: joining, resolving, and separating sister chromatids during mitosis and meiosis. Annu. Rev. Genet. 35, 673–745 (2001).

    Article  CAS  Google Scholar 

  33. Pradhan, B. et al. The Smc5/6 complex is a DNA loop extruding motor. Preprint at bioRxiv https://doi.org/10.1101/2022.05.13.491800 (2022).

  34. Venegas, A. B., Natsume, T., Kanemaki, M. & Hickson, I. D. Inducible degradation of the human SMC5/6 complex reveals an essential role only during interphase. Cell Rep. 31, 107533 (2020).

    Article  CAS  Google Scholar 

  35. Zabolotnaya, E., Mela, I., Henderson, R. M. & Robinson, N. P. Turning the Mre11/Rad50 DNA repair complex on its head: lessons from SMC protein hinges, dynamic coiled-coil movements and DNA loop-extrusion? Biochem. Soc. Trans. 48, 2359–2376 (2020).

    Article  CAS  Google Scholar 

  36. Gurzau, A. D., Blewitt, M. E., Czabotar, P. E., Murphy, J. M. & Birkinshaw, R. W. Relating SMCHD1 structure to its function in epigenetic silencing. Biochem. Soc. Trans. 48, 1751–1763 (2020).

    Article  CAS  Google Scholar 

  37. Waizenegger, I. C., Hauf, S., Meinke, A. & Peters, J.-M. Two distinct pathways remove mammalian cohesin from chromosome arms in prophase and from centromeres in anaphase. Cell 103, 399–410 (2000).

    Article  CAS  Google Scholar 

  38. Goloborodko, A., Imakaev, M. V., Marko, J. F. & Mirny, L. Compaction and segregation of sister chromatids via active loop extrusion. eLife 5, e14864 (2016).

    Article  Google Scholar 

  39. Houlard, M. et al. MCPH1 inhibits condensin II during interphase by regulating its SMC2–kleisin interface. eLife 10, e73348 (2021).

    Article  CAS  Google Scholar 

  40. Oldenkamp, R. & Rowland, B. D. A walk through the SMC cycle: from catching DNAs to shaping the genome. Mol. Cell 82, 1616–1630 (2022).

    Article  CAS  Google Scholar 

  41. Elbatsh, A. M. O. et al. Distinct roles for condensin’s two ATPase sites in chromosome condensation. Mol. Cell 76, 724–737.e5 (2019).

    Article  CAS  Google Scholar 

  42. Perea-Resa, C., Wattendorf, L., Marzouk, S. & Blower, M. D. Cohesin: behind dynamic genome topology and gene expression reprogramming. Trends Cell Biol. 31, 760–773 (2021).

    Article  CAS  Google Scholar 

  43. Petela, N. J. et al. Scc2 is a potent activator of cohesin’s ATPase that promotes loading by binding Scc1 without Pds5. Mol. Cell 70, 1134–1148.e7 (2018).

    Article  CAS  Google Scholar 

  44. Kikuchi, S., Borek, D. M., Otwinowski, Z., Tomchick, D. R. & Yu, H. Crystal structure of the cohesin loader Scc2 and insight into cohesinopathy. Proc. Natl Acad. Sci. USA 113, 12444–12449 (2016).

    Article  CAS  Google Scholar 

  45. Dauban, L. et al. Regulation of cohesin-mediated chromosome folding by Eco1 and other partners. Mol. Cell 77, 1279–1293.e4 (2020).

    Article  CAS  Google Scholar 

  46. Wutz, G. et al. ESCO1 and CTCF enable formation of long chromatin loops by protecting cohesinSTAG1 from WAPL. eLife 9, e52091 (2020). This work shows that Hi-C after acute depletion of cohesin, WAPL/PDS5 and CTCF contrasts their relative contribution to genome folding.

    Article  CAS  Google Scholar 

  47. van Ruiten, M. S. et al. The cohesin acetylation cycle controls chromatin loop length through a PDS5A brake mechanism. Nat. Struct. Mol. Biol. 29, 586–591 (2022).

    Article  Google Scholar 

  48. Bastié, N. et al. Smc3 acetylation, Pds5 and Scc2 control the translocase activity that establishes cohesin-dependent chromatin loops. Nat. Struct. Mol. Biol. 29, 575–585 (2022).

    Article  Google Scholar 

  49. van Ruiten, M. S. & Rowland, B. D. SMC complexes: universal DNA looping machines with distinct regulators. Trend Genet. 34, 477–487 (2018).

    Article  Google Scholar 

  50. Haarhuis, J. H. I. et al. The cohesin release factor WAPL restricts chromatin loop extension. Cell 169, 693–707.e14 (2017).

    Article  CAS  Google Scholar 

  51. 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).

    Article  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

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

    Article  Google Scholar 

  54. 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).

    Article  CAS  Google Scholar 

  55. Vietri Rudan, M. et al. Comparative Hi-C reveals that CTCF underlies evolution of chromosomal domain architecture. Cell Rep. 10, 1297–1309 (2015).

    Article  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

  57. Nora, E. P. et al. Molecular basis of CTCF binding polarity in genome folding. Nat. Commun. 11, 5612 (2020). This paper performs separation of function analysis of different protein domains by complementing inducible degradation of endogenous CTCF with mutants to show the importance of the N-terminal domain in CTCF-anchored loop formation.

    Article  CAS  Google Scholar 

  58. Davidson, I. F. et al. DNA loop extrusion by human cohesin. Science 366, 1338–1345 (2019).

    Article  CAS  Google Scholar 

  59. Kim, Y., Shi, Z., Zhang, H., Finkelstein, I. J. & Yu, H. Human cohesin compacts DNA by loop extrusion. Science 366, 1345–1349 (2019).

    Article  CAS  Google Scholar 

  60. Gabriele, M. et al. Dynamics of CTCF- and cohesin-mediated chromatin looping revealed by live-cell imaging. Science 376, 496–501 (2022).

    Article  CAS  Google Scholar 

  61. Mach, P. et al. Live-cell imaging and physical modeling reveal control of chromosome folding dynamics by cohesin and CTCF. Preprint at bioRxiv https://doi.org/10.1101/2022.03.03.482826 (2022).

  62. Trauth, J., Scheffer, J., Hasenjäger, S. & Taxis, C. Synthetic control of protein degradation during cell proliferation and developmental processes. ACS Omega 4, 2766–2778 (2019).

    Article  CAS  Google Scholar 

  63. Verma, R., Mohl, D. & Deshaies, R. J. Harnessing the power of proteolysis for targeted protein inactivation. Mol. Cell 77, 446–460 (2020).

    Article  CAS  Google Scholar 

  64. Kanemaki, M. T. Ligand-induced degrons for studying nuclear functions. Curr. Opin. Cell Biol. 74, 29–36 (2022).

    Article  CAS  Google Scholar 

  65. Nishimura, K., Fukagawa, T., Takisawa, H., Kakimoto, T. & Kanemaki, M. An auxin-based degron system for the rapid depletion of proteins in nonplant cells. Nat. Methods 6, 917–922 (2009). This paper describes the first use of the AID system for targeted protein depletion outside plants.

    Article  CAS  Google Scholar 

  66. Yesbolatova, A. et al. The auxin-inducible degron 2 technology provides sharp degradation control in yeast, mammalian cells, and mice. Nat. Commun. 11, 5701 (2020).

    Article  CAS  Google Scholar 

  67. Li, S., Prasanna, X., Salo, V. T., Vattulainen, I. & Ikonen, E. An efficient auxin-inducible degron system with low basal degradation in human cells. Nat. Methods 16, 866–869 (2019).

    Article  CAS  Google Scholar 

  68. Nabet, B. et al. The dTAG system for immediate and target-specific protein degradation. Nat. Chem. Biol. 14, 431–441 (2018). This paper establishes the FKBP12/dTAG system.

    Article  CAS  Google Scholar 

  69. Nabet, B. et al. Rapid and direct control of target protein levels with VHL-recruiting dTAG molecules. Nat. Commun. 11, 4687 (2020).

    Article  CAS  Google Scholar 

  70. Tovell, H. et al. Rapid and reversible knockdown of endogenously tagged endosomal proteins via an optimized HaloPROTAC degrader. ACS Chem. Biol. 14, 882–892 (2019).

    Article  CAS  Google Scholar 

  71. Bond, A. G. et al. Development of BromoTag: a ‘bump-and-hole’-PROTAC system to induce potent, rapid, and selective degradation of tagged target proteins. J. Med. Chem. 64, 15477–15502 (2021).

    Article  CAS  Google Scholar 

  72. Nowak, R. P. et al. Structure-guided design of a ‘bump-and-hole’ bromodomain-based degradation tag. J. Med. Chem. 64, 11637–11650 (2021).

    Article  CAS  Google Scholar 

  73. Békés, M., Langley, D. R. & Crews, C. M. PROTAC targeted protein degraders: the past is prologue. Nat. Rev. Drug Discov. 21, 181–200 (2022).

    Article  Google Scholar 

  74. Röth, S., Fulcher, L. J. & Sapkota, G. P. Advances in targeted degradation of endogenous proteins. Cell Mol. Life Sci. 76, 2761–2777 (2019).

    Article  Google Scholar 

  75. Usherenko, S. et al. Photo-sensitive degron variants for tuning protein stability by light. BMC Syst. Biol. 8, 128 (2014).

    Article  Google Scholar 

  76. Langan, R. A. et al. De novo design of bioactive protein switches. Nature 572, 205–210 (2019).

    Article  CAS  Google Scholar 

  77. Samejima, K. et al. Functional analysis after rapid degradation of condensins and 3D-EM reveals chromatin volume is uncoupled from chromosome architecture in mitosis. J. Cell Sci. 131, jcs210187 (2018).

    Article  Google Scholar 

  78. Hoencamp, C. et al. 3D genomics across the tree of life reveals condensin II as a determinant of architecture type. Science 372, 984–989 (2021).

    Article  CAS  Google Scholar 

  79. Strom, A. R. et al. HP1α is a chromatin crosslinker that controls nuclear and mitotic chromosome mechanics. eLife 10, e63972 (2021).

    Article  CAS  Google Scholar 

  80. Meijering, A. E. C. et al. Nonlinear mechanics of human mitotic chromosomes. Nature 605, 545–550 (2022).

    Article  CAS  Google Scholar 

  81. Takagi, M. et al. Ki-67 and condensins support the integrity of mitotic chromosomes through distinct mechanisms. J. Cell Sci. 131, jcs212092 (2018).

    Article  Google Scholar 

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

    Article  CAS  Google Scholar 

  83. Rao, S. S. P. et al. Cohesin loss eliminates all loop domains. Cell 171, 305–320.e24 (2017). This work shows that acute depletion of cohesin reveals the role of cohesin in TAD and loop formation genome-wide. Restoration of cohesin following depletion shows that structures are reformed within a few hours, highlighting the dynamics of the system.

    Article  CAS  Google Scholar 

  84. Thiecke, M. J. et al. Cohesin-dependent and -independent mechanisms mediate chromosomal contacts between promoters and enhancers. Cell Rep. 32, 107929 (2020).

    Article  CAS  Google Scholar 

  85. Rhodes, J. D. P. et al. Cohesin disrupts Polycomb-dependent chromosome interactions in embryonic stem cells. Cell Rep. 30, 820–835.e10 (2020).

    Article  CAS  Google Scholar 

  86. Luppino, J. M. et al. Cohesin promotes stochastic domain intermingling to ensure proper regulation of boundary-proximal genes. Nat. Genet. 52, 840–848 (2020).

    Article  CAS  Google Scholar 

  87. Szabo, Q. et al. Regulation of single-cell genome organization into TADs and chromatin nanodomains. Nat. Genet. 52, 1151–1157 (2020).

    Article  CAS  Google Scholar 

  88. Hafner, A., Park, M., Berger, S. E., Nora, E. P. & Boettiger, A. N. Beyond TADs: microscopy reveals 2 axes of chromosome organization by CTCF and cohesin. Preprint at bioRxiv https://doi.org/10.1101/2022.07.13.499982 (2022).

  89. Hsieh, T.-H. S. et al. Enhancer–promoter interactions and transcription are maintained upon acute loss of CTCF, cohesin, WAPL, and YY1. Preprint at bioRxiv https://doi.org/10.1101/2021.07.14.452365 (2021). This preprint provides preliminary evidence that acute cohesin of CTCF, cohesin, WAPL or YY1 only has a minor immediate effect on enhancer–promoter interactions as detected by Micro-C.

  90. Aljahani, A. et al. Analysis of sub-kilobase chromatin topology reveals nano-scale regulatory interactions with variable dependence on cohesin and CTCF. Nat. Commun. 13, 2139 (2022).

    Article  CAS  Google Scholar 

  91. Goel, V. Y., Huseyin, M. K. & Hansen, A. S. Region capture Micro-C reveals coalescence of enhancers and promoters into nested microcompartments. Preprint at bioRxiv https://doi.org/10.1101/2022.07.12.499637 (2022).

  92. Joshi, O. et al. Dynamic reorganization of extremely long-range promoter–promoter interactions between two states of pluripotency. Cell Stem Cell 17, 748–757 (2015).

    Article  CAS  Google Scholar 

  93. 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–E6706 (2018).

    Article  CAS  Google Scholar 

  94. Zhu, Y., Denholtz, M., Lu, H. & Murre, C. Calcium signaling instructs NIPBL recruitment at active enhancers and promoters via distinct mechanisms to reconstruct genome compartmentalization. Genes Dev. 35, 65–81 (2021).

    Article  CAS  Google Scholar 

  95. Kriz, A. J., Colognori, D., Sunwoo, H., Nabet, B. & Lee, J. T. Balancing cohesin eviction and retention prevents aberrant chromosomal interactions, Polycomb-mediated repression, and X-inactivation. Mol. Cell 81, 1970–1987.e9 (2021).

    Article  CAS  Google Scholar 

  96. Liu, N. Q. et al. WAPL maintains a cohesin loading cycle to preserve cell-type-specific distal gene regulation. Nat. Genet. 53, 100–109 (2021). This study reveals the crucial role of the cohesin loading cycle for maintaining lineage-specific gene expression.

    Article  CAS  Google Scholar 

  97. Liu, N. Q. et al. Rapid depletion of CTCF and cohesin proteins reveals dynamic features of chromosome architecture. Preprint at bioRxiv https://doi.org/10.1101/2021.08.27.457977 (2021).

  98. Miron, E. et al. Chromatin arranges in chains of mesoscale domains with nanoscale functional topography independent of cohesin. Sci. Adv. 6, eaba8811 (2020).

    Article  CAS  Google Scholar 

  99. van Schaik, T. et al. CTCF and cohesin promote focal detachment of DNA from the nuclear lamina. Genome Biol. 23, 185 (2022).

    Article  Google Scholar 

  100. Ladurner, R. et al. Sororin actively maintains sister chromatid cohesion. EMBO J. 35, 635–653 (2016).

    Article  CAS  Google Scholar 

  101. Mitter, M. et al. Conformation of sister chromatids in the replicated human genome. Nature 586, 139–144 (2020).

    Article  CAS  Google Scholar 

  102. Casa, V. et al. Redundant and specific roles of cohesin STAG subunits in chromatin looping and transcriptional control. Genome Res. 30, 515–527 (2020).

    Article  CAS  Google Scholar 

  103. Cuadrado, A. et al. Specific contributions of cohesin–SA1 and cohesin–SA2 to TADs and polycomb domains in embryonic stem cells. Cell Rep. 27, 3500–3510.e4 (2019).

    Article  CAS  Google Scholar 

  104. Kojic, A. et al. Distinct roles of cohesin–SA1 and cohesin–SA2 in 3D chromosome organization. Nat. Struct. Mol. Biol. 25, 496–504 (2018).

    Article  CAS  Google Scholar 

  105. van der Weide, R. H. et al. Hi-C analyses with GENOVA: a case study with cohesin variants. NAR Genom. Bioinform 3, lqab040 (2021).

    Article  Google Scholar 

  106. Romero-Pérez, L., Surdez, D., Brunet, E., Delattre, O. & Grünewald, T. G. P. STAG mutations in cancer. Trends Cancer 5, 506–520 (2019).

    Article  Google Scholar 

  107. 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).

    Article  CAS  Google Scholar 

  108. Splinter, E. CTCF mediates long-range chromatin looping and local histone modification in the β-globin locus. Genes Dev. 20, 2349–2354 (2006).

    Article  CAS  Google Scholar 

  109. Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944.e22 (2017). Hi-C following acute depletion of CTCF shows that CTCF acts as a boundary factor genome-wide.

    Article  CAS  Google Scholar 

  110. Nishana, M. et al. Defining the relative and combined contribution of CTCF and CTCFL to genomic regulation. Genome Biol. 21, 108 (2020).

    Article  CAS  Google Scholar 

  111. Pugacheva, E. M. et al. CTCF mediates chromatin looping via N-terminal domain-dependent cohesin retention. Proc. Natl Acad. Sci. USA 117, 2020–2031 (2020).

    Article  CAS  Google Scholar 

  112. Saldana-Meyer, R. et al. RNA interactions with CTCF are essential for its proper function. Mol. Cell 76, 412–422. (2019).

    CAS  Google Scholar 

  113. 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.e13 (2019).

    Article  CAS  Google Scholar 

  114. Luan, J. et al. CTCF blocks anti-sense transcription initiation at divergent gene promoters. Preprint at bioRxiv https://doi.org/10.1101/2021.10.30.465508 (2021).

  115. Linares-Saldana, R. et al. BRD4 orchestrates genome folding to promote neural crest differentiation. Nat. Genet. 53, 1480–1492 (2021).

    Article  CAS  Google Scholar 

  116. Crump, N. T. et al. BET inhibition disrupts transcription but retains enhancer–promoter contact. Nat. Commun. 12, 223 (2021).

    Article  CAS  Google Scholar 

  117. 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.e20 (2019).

    Article  CAS  Google Scholar 

  118. Ramasamy, S. et al. The Mediator complex regulates enhancer–promoter interactions. Preprint at bioRxiv https://doi.org/10.1101/2022.06.15.496245 (2022).

  119. Haarhuis, J. H. I. et al. A Mediator–cohesin axis controls heterochromatin domain formation. Nat. Commun. 13, 754 (2022).

    Article  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

  121. Spracklin, G. et al. Heterochromatin diversity modulates genome compartmentalization and loop extrusion barriers. Preprint at bioRxiv https://doi.org/10.1101/2021.08.05.455340 (2021).

  122. Dequeker, B. J. H. et al. MCM complexes are barriers that restrict cohesin-mediated loop extrusion. Nature 606, 197–203 (2022).

    Article  CAS  Google Scholar 

  123. Banigan, E. J. et al. Transcription shapes 3D chromatin organization by interacting with loop-extruding cohesin complexes. Preprint at bioRxiv https://doi.org/10.1101/2022.01.07.475367 (2022).

  124. Zhang, S., Übelmesser, N., Barbieri, M. & Papantonis, A. Enhancer–promoter contact formation requires RNAPII and antagonizes loop extrusion. Preprint at bioRxiv https://doi.org/10.1101/2022.07.04.498738 (2022).

  125. Jiang, Y. et al. Genome-wide analyses of chromatin interactions after the loss of Pol I, Pol II, and Pol III. Genome Biol. 21, 158 (2020).

    Article  CAS  Google Scholar 

  126. Zhang, S. et al. RNA polymerase II is required for spatial chromatin reorganization following exit from mitosis. Sci. Adv. 7, eabg8205 (2021). Together with Jiang et al. (2020), this work uses RNA polymerase degrons to show the limited role of these complexes in 3D genome organization, as assayed by Hi-C.

    Article  CAS  Google Scholar 

  127. Neguembor, M. V. et al. Transcription-mediated supercoiling regulates genome folding and loop formation. Mol. Cell 81, 3065–3081.e12 (2021).

    Article  CAS  Google Scholar 

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

    Article  CAS  Google Scholar 

  129. 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).

    Article  CAS  Google Scholar 

  130. Kubo, N. et al. Promoter-proximal CTCF binding promotes distal enhancer-dependent gene activation. Nat. Struct. Mol. Biol. 28, 152–161 (2021).

    Article  CAS  Google Scholar 

  131. Ottema, S. et al. The leukemic oncogene EVI1 hijacks a MYC super-enhancer by CTCF-facilitated loops. Nat. Commun. 12, 5679 (2021).

    Article  CAS  Google Scholar 

  132. Schuijers, J. et al. Transcriptional dysregulation of MYC reveals common enhancer-docking mechanism. Cell Rep. 23, 349–360 (2018).

    Article  CAS  Google Scholar 

  133. Benabdallah, N. S. et al. Decreased enhancer–promoter proximity accompanying enhancer activation. Mol. Cell 76, 473–484.e7 (2019).

    Article  CAS  Google Scholar 

  134. Karr, J. P., Ferrie, J. J., Tjian, R. & Darzacq, X. The transcription factor activity gradient (TAG) model: contemplating a contact-independent mechanism for enhancer–promoter communication. Genes Dev. 36, 7–16 (2022).

    Article  CAS  Google Scholar 

  135. Li, J. & Pertsinidis, A. New insights into promoter–enhancer communication mechanisms revealed by dynamic single-molecule imaging. Biochem. Soc. Trans. 49, 1299–1309 (2021).

    Article  CAS  Google Scholar 

  136. Stelloh, C. et al. The cohesin-associated protein Wapal is required for proper Polycomb-mediated gene silencing. Epigenetics Chromatin 9, 14 (2016).

    Article  Google Scholar 

  137. Luppino, J. M. et al. NIPBL and WAPL balance cohesin activity to regulate chromatin folding and gene expression. Preprint at bioRxiv https://doi.org/10.1101/2022.04.19.488785 (2022).

  138. Kean, C. M. et al. Decreasing Wapl dosage partially corrects transcriptome phenotypes in Nipbl–/+ embryonic mouse brain. Preprint at bioRxiv https://doi.org/10.1101/2022.05.31.493745 (2022).

  139. Zuin, J. et al. Nonlinear control of transcription through enhancer-promoter interactions. Nature 604, 571–577 (2022).

    Article  CAS  Google Scholar 

  140. Xie, L. et al. BRD2 compartmentalizes the accessible genome. Nat. Genet. 54, 481–491 (2022).

    Article  CAS  Google Scholar 

  141. Rinaldi, L. et al. The glucocorticoid receptor associates with the cohesin loader NIPBL to promote long-range gene regulation. Sci. Adv. 8, eabj8360 (2022).

    Article  CAS  Google Scholar 

  142. Gu, B. et al. Opposing effects of cohesin and transcription on CTCF organization revealed by super-resolution imaging. Mol. Cell 80, 699–711.e7 (2020).

    Article  CAS  Google Scholar 

  143. Lee, R. et al. CTCF-mediated chromatin looping provides a topological framework for the formation of phase-separated transcriptional condensates. Nucleic Acids Res. 50, 207–226 (2022).

    Article  CAS  Google Scholar 

  144. Nozaki, T. et al. Dynamic organization of chromatin domains revealed by super-resolution live-cell imaging. Mol. Cell 67, 282–293.e7 (2017).

    Article  CAS  Google Scholar 

  145. Zuin, J. et al. Nonlinear control of transcription through enhancer–promoter interactions. Nature 604, 571–577 (2022).

    Article  CAS  Google Scholar 

  146. Rinzema, N. J. et al. Building regulatory landscapes reveals that an enhancer can recruit cohesin to create contact domains, engage CTCF sites and activate distant genes. Nat. Struct. Mol. Biol. 29, 563–574 (2022).

    Article  CAS  Google Scholar 

  147. Kane, L. et al. Cohesin is required for long-range enhancer action. Preprint at bioRxiv https://doi.org/10.1101/2021.06.24.449812 (2021).

  148. Weintraub, A. S. et al. YY1 is a structural regulator of enhancer–promoter loops. Cell 171, 1573–1588.e28 (2017).

    Article  CAS  Google Scholar 

  149. Dobrinić, P., Szczurek, A. T. & Klose, R. J. PRC1 drives Polycomb-mediated gene repression by controlling transcription initiation and burst frequency. Nat. Struct. Mol. Biol. 28, 811–824 (2021).

    Article  Google Scholar 

  150. Barshad, G. et al. RNA polymerase II and PARP1 shape enhancer-promoter contacts. Preprint at bioRxiv https://doi.org/10.1101/2022.07.07.499190 (2022).

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

    Article  CAS  Google Scholar 

  152. Saldaña-Meyer, R. et al. RNA interactions are essential for CTCF-mediated genome organization. Mol. Cell 76, 412–422.e5 (2019).

    Article  Google Scholar 

  153. Stik, G. et al. CTCF is dispensable for immune cell transdifferentiation but facilitates an acute inflammatory response. Nat. Genet. 52, 655–661 (2020).

    Article  CAS  Google Scholar 

  154. Pezic, D. et al. The cohesin regulator Stag1 promotes cell plasticity through heterochromatin regulation. Preprint at bioRxiv https://doi.org/10.1101/2021.02.14.429938 (2021).

  155. Zhang, K. et al. Analysis of genome architecture during SCNT reveals a role of cohesin in impeding minor ZGA. Mol. Cell 79, 234–250.e9 (2020).

    Article  CAS  Google Scholar 

  156. Zhu, Y. et al. Relaxed 3D genome conformation facilitates the pluripotent to totipotent-like state transition in embryonic stem cells. Nucleic Acids Res. 49, 12167–12177 (2021).

    Article  CAS  Google Scholar 

  157. Olbrich, T. et al. CTCF is a barrier for 2C-like reprogramming. Nat. Commun. 12, 4856 (2021).

    Article  CAS  Google Scholar 

  158. Flavahan, W. A. et al. Altered chromosomal topology drives oncogenic programs in SDH-deficient GISTs. Nature 575, 229–233 (2019).

    Article  CAS  Google Scholar 

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

    Article  Google Scholar 

  160. Valton, A.-L. & Dekker, J. TAD disruption as oncogenic driver. Curr. Opin. Genet. Dev. 36, 34–40 (2016).

    Article  CAS  Google Scholar 

  161. Xiao, J. Y., Hafner, A. & Boettiger, A. N. How subtle changes in 3D structure can create large changes in transcription. eLife 10, e64320 (2021).

    Article  CAS  Google Scholar 

  162. Alexander, J. M. et al. Live-cell imaging reveals enhancer-dependent Sox2 transcription in the absence of enhancer proximity. eLife 8, e41769 (2019).

    Article  Google Scholar 

  163. Jerkovic, I. & Cavalli, G. Understanding 3D genome organization by multidisciplinary methods. Nat. Rev. Mol. Cell Biol. 22, 511–528 (2021).

    Article  CAS  Google Scholar 

  164. Kline, A. D. et al. Diagnosis and management of Cornelia de Lange syndrome: first international consensus statement. Nat. Rev. Genet. 19, 649–666 (2018).

    Article  CAS  Google Scholar 

  165. Solomon, D. A., Kim, J.-S. & Waldman, T. Cohesin gene mutations in tumorigenesis: from discovery to clinical significance. BMB Rep. 47, 299–310 (2014).

    Article  Google Scholar 

  166. Kiefer, L. et al. Cohesin erases genomic-proximity biases to drive stochastic protocadherin expression for proper neural wiring. Preprint at bioRxiv https://doi.org/10.1101/2022.03.09.483674 (2022).

  167. Canzio, D. et al. Antisense lncRNA transcription mediates DNA demethylation to drive stochastic protocadherin α promoter choice. Cell 177, 639–653.e15 (2019).

    Article  CAS  Google Scholar 

  168. Ba, Z. et al. CTCF orchestrates long-range cohesin-driven V(D)J recombinational scanning. Nature 586, 305–331 (2020).

    Article  CAS  Google Scholar 

  169. Vian, L. et al. The energetics and physiological impact of cohesin extrusion. Cell 173, 1165–1178.e20 (2018).

    Article  CAS  Google Scholar 

  170. Zhang, X. et al. Fundamental roles of chromatin loop extrusion in antibody class switching. Nature 575, 385–389 (2019).

    Article  CAS  Google Scholar 

  171. Zhang, Y. et al. The fundamental role of chromatin loop extrusion in physiological V(D)J recombination. Nature 573, 600–604 (2019).

    Article  CAS  Google Scholar 

  172. Dai, H.-Q. et al. Loop extrusion mediates physiological Igh locus contraction for RAG scanning. Nature 590, 338–343 (2021).

    Article  CAS  Google Scholar 

  173. Hill, L. et al. Wapl repression by Pax5 promotes V gene recombination by Igh loop extrusion. Nature 584, 142–147 (2020).

    Article  CAS  Google Scholar 

  174. Arnould, C. et al. Loop extrusion as a mechanism for formation of DNA damage repair foci. Nature 590, 660–665 (2021).

    Article  CAS  Google Scholar 

  175. Yunusova, A. et al. Evaluation of the OsTIR1 and AtAFB2 AID systems for genome architectural protein degradation in mammalian cells. Front. Mol. Biosci. 8, 757394 (2021).

    Article  CAS  Google Scholar 

  176. Naqvi, S. et al. Precise modulation of transcription factor levels reveals drivers of dosage sensitivity. Preprint at bioRxiv https://doi.org/10.1101/2022.06.13.495964 (2022).

  177. Zhang, H. et al. CTCF and transcription influence chromatin structure re-configuration after mitosis. Nat. Commun. 12, 5157 (2021).

    Article  CAS  Google Scholar 

  178. Capece, M. et al. A novel auxin-inducible degron system for rapid, cell cycle-specific targeted proteolysis. Preprint at bioRxiv https://doi.org/10.1101/2021.04.23.441203 (2021).

  179. Pryzhkova, M. V., Xu, M. J. & Jordan, P. W. Adaptation of the AID system for stem cell and transgenic mouse research. Stem Cell Res. 49, 102078 (2020).

    Article  CAS  Google Scholar 

  180. Macdonald, L. et al. Rapid and specific degradation of endogenous proteins in mouse models using auxin-inducible degrons. eLife 11, e77987 (2022).

    Article  CAS  Google Scholar 

  181. Zhang, L., Ward, J. D., Cheng, Z. & Dernburg, A. F. The auxin-inducible degradation (AID) system enables versatile conditional protein depletion in C. elegans. Development 142, 4374–4384 (2015).

    CAS  Google Scholar 

  182. Abuhashem, A., Lee, A. S., Joyner, A. L. & Hadjantonakis, A.-K. Rapid and efficient degradation of endogenous proteins in vivo identifies stage-specific roles of RNA Pol II pausing in mammalian development. Dev. Cell 57, 1068–1080.e6 (2022).

    Article  CAS  Google Scholar 

  183. Luan, J. et al. Distinct properties and functions of CTCF revealed by a rapidly inducible degron system. Cell Rep. 34, 108783 (2021).

    Article  CAS  Google Scholar 

Download references

Acknowledgements

The authors are grateful to L. Braccioli and R. Shah for feedback on the manuscript. E.d.W. is supported by a European Research Council (ERC) Consolidator grant FuncDis3D (865459) and is part of Oncode which is partly financed by the Dutch Cancer Society. E.P.N. is supported by National Institutes of Health (NIH) grant 1R35GM142792-01.

Author information

Authors and Affiliations

  1. Division of Gene Regulation, Oncode Institute, Amsterdam, the Netherlands

    Elzo de Wit

  2. Division of Gene Regulation, Netherlands Cancer Institute, Amsterdam, the Netherlands

    Elzo de Wit

  3. Cardiovascular Research Institute, University of California, San Francisco, San Francisco, CA, USA

    Elphège P. Nora

  4. Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA, USA

    Elphège P. Nora

Authors
  1. Elzo de Wit
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Elphège P. Nora
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

The authors contributed equally to all aspects of the article.

Corresponding authors

Correspondence to Elzo de Wit or Elphège P. Nora.

Ethics declarations

Competing interests

The authors declare no competing interests.

Peer review

Peer review information

Nature Reviews Genetics thanks the anonymous reviewers for their contribution to the peer review of this work.

Additional information

Publisher’s note

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

Glossary

CTCF

An 11 zinc finger DNA and RNA-binding protein able to block the extrusion of cohesin in part through its amino-terminal domain that interacts with the STAG and RAD21 cohesin subunits, thereby shielding it from WAPL-mediated release.

Topologically associating domains

(TADs). Genomic regions that show preferential self-interaction and are physically insulated from their neighbours by the binding of CTCF at boundaries. Self-interaction is mediated by the cohesin complex.

Super-enhancers

Multi-kilobase stretches of regulatory DNA that exhibit unusually strong occupancy of transcription factors and co-factors.

Polycomb

A family of chromatin proteins that assemble into the (generally) repressive complexes Polycomb repressive complex 1 (PRC1) and PRC2, which maintain epigenetic silencing of developmental genes.

Cohesive cohesin

A pool of cohesin complex that mediates sister chromatin cohesion, which depends on structural maintenance of chromosomes 3 (SMC3) acetylation, PDS5 and SORORIN binding. Cohesive cohesin is thought to be distinct from extrusive cohesin as it contains PDS5 (and not NIPBL).

Phase separation

A process through which polymer chains (or segments of a polymer chain) spontaneously de-mix and segregate through the formation of immiscible phases.

Rights and permissions

Springer Nature or its licensor holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

de Wit, E., Nora, E.P. New insights into genome folding by loop extrusion from inducible degron technologies. Nat Rev Genet 24, 73–85 (2023). https://doi.org/10.1038/s41576-022-00530-4

Download citation

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41576-022-00530-4

This article is cited by

Search

Advanced search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing