Abstract
The primary regulators of metazoan gene expression are enhancers, originally functionally defined as DNA sequences that can activate transcription at promoters in an orientation-independent and distance-independent manner. Despite being crucial for gene regulation in animals, what mechanisms underlie enhancer selectivity for promoters, and more fundamentally, how enhancers interact with promoters and activate transcription, remain poorly understood. In this Review, we first discuss current models of enhancer–promoter interactions in space and time and how enhancers affect transcription activation. Next, we discuss different mechanisms that mediate enhancer selectivity, including repression, biochemical compatibility and regulation of 3D genome structure. Through 3D polymer simulations, we illustrate how the ability of 3D genome folding mechanisms to mediate enhancer selectivity strongly varies for different enhancer–promoter interaction mechanisms. Finally, we discuss how recent technical advances may provide new insights into mechanisms of enhancer–promoter interactions and how technical biases in methods such as Hi-C and Micro-C and imaging techniques may affect their interpretation.
This is a preview of subscription content, access via your institution
Access options
Access Nature and 54 other Nature Portfolio journals
Get Nature+, our best-value online-access subscription
$29.99 / 30 days
cancel any time
Subscribe to this journal
Receive 12 print issues and online access
$189.00 per year
only $15.75 per issue
Buy this article
- Purchase on Springer Link
- Instant access to full article PDF
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
References
Bentovim, L., Harden, T. T. & DePace, A. H. Transcriptional precision and accuracy in development: from measurements to models and mechanisms. Development 144, 3855–3866 (2017).
Ong, C.-T. & Corces, V. G. Enhancer function: new insights into the regulation of tissue-specific gene expression. Nat. Rev. Genet. 12, 283–293 (2011).
Field, A. & Adelman, K. Evaluating enhancer function and transcription. Annu. Rev. Biochem. 89, 213–234 (2020).
Zabidi, M. A. & Stark, A. Regulatory enhancer–core-promoter communication via transcription factors and cofactors. Trends Genet. 32, 801–814 (2016).
Andersson, R. et al. An atlas of active enhancers across human cell types and tissues. Nature 507, 455–461 (2014).
Spitz, F. & Furlong, E. E. Transcription factors: from enhancer binding to developmental control. Nat. Rev. Genet. 13, 613–626 (2012).
Banerji, J., Olson, L. & Schaffner, W. A lymphocyte-specific cellular enhancer is located downstream of the joining region in immunoglobulin heavy chain genes. Cell 33, 729–740 (1983).
Gillies, S. D., Morrison, S. L., Oi, V. T. & Tonegawa, S. A tissue-specific transcription enhancer element is located in the major intron of a rearranged immunoglobulin heavy chain gene. Cell 33, 717–728 (1983).
Mercola, M., Wang, X.-F., Olsen, J. & Calame, K. Transcriptional enhancer elements in the mouse immunoglobulin heavy chain locus. Science 221, 663–665 (1983).
Banerji, J., Rusconi, S. & Schaffner, W. Expression of a β-globin gene is enhanced by remote SV40 DNA sequences. Cell 27, 299–308 (1981).
Halfon, M. S. Studying transcriptional enhancers: the founder fallacy, validation creep, and other biases. Trends Genet. 35, 93–103 (2019).
Galouzis, C. C. & Furlong, E. E. Regulating specificity in enhancer–promoter communication. Curr. Opin. Cell Biol. 75, 102065 (2022).
van Arensbergen, J., van Steensel, B. & Bussemaker, H. J. In search of the determinants of enhancer–promoter interaction specificity. Trends Cell Biol. 24, 695–702 (2014).
Furlong, E. E. & Levine, M. Developmental enhancers and chromosome topology. Science 361, 1341–1345 (2018).
Moreau, P. et al. The SV40 72 base repair repeat has a striking effect on gene expression both in SV40 and other chimeric recombinants. Nucleic Acids Res. 9, 6047–6068 (1981).
Travers, A. Chromatin modification by DNA tracking. Proc. Natl Acad. Sci. USA 96, 13634–13637 (1999).
Hatzis, P. & Talianidis, I. Dynamics of enhancer–promoter communication during differentiation-induced gene activation. Mol. Cell 10, 1467–1477 (2002).
Bulger, M. & Groudine, M. Looping versus linking: toward a model for long-distance gene activation. Genes Dev. 13, 2465–2477 (1999).
Chen, Z. et al. Widespread increase in enhancer–promoter interactions during developmental enhancer activation in mammals. Preprint at bioRxiv https://doi.org/10.1101/2022.11.18.516017 (2022).
Gasperini, M. et al. A genome-wide framework for mapping gene regulation via cellular genetic screens. Cell 176, 377–390 (2019).
Li, G. et al. Extensive promoter-centered chromatin interactions provide a topological basis for transcription regulation. Cell 148, 84–98 (2012).
Sanyal, A., Lajoie, B. R., Jain, G. & Dekker, J. The long-range interaction landscape of gene promoters. Nature 489, 109–113 (2012).
Goel, V. Y., Huseyin, M. K. & Hansen, A. S. Region capture Micro-C reveals coalescence of enhancers and promoters into nested microcompartments. Nat. Genet. 55, 1048–1056 (2023).
Zuin, J. et al. Nonlinear control of transcription through enhancer–promoter interactions. Nature 604, 571–577 (2022).
Brückner, D. B., Chen, H., Barinov, L., Zoller, B. & Gregor, T. Stochastic motion and transcriptional dynamics of pairs of distal DNA loci on a compacted chromosome. Science 380, 1357–1362 (2023).
Mateo, L. J. et al. Visualizing DNA folding and RNA in embryos at single-cell resolution. Nature 568, 49–54 (2019).
Chen, H. et al. Dynamic interplay between enhancer–promoter topology and gene activity. Nat. Genet. 50, 1296–1303 (2018).
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).
Deng, W. et al. Reactivation of developmentally silenced globin genes by forced chromatin looping. Cell 158, 849–860 (2014).
Hsieh, T.-H. S. et al. Resolving the 3D landscape of transcription-linked mammalian chromatin folding. Mol. Cell 78, 539–553 (2020).
Hsieh, T.-H. S. et al. Enhancer–promoter interactions and transcription are largely maintained upon acute loss of CTCF, cohesin, WAPL or YY1. Nat. Genet. 54, 1919–1932 (2022).
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).
Fulco, C. P. et al. Activity-by-contact model of enhancer–promoter regulation from thousands of CRISPR perturbations. Nat. Genet. 51, 1664–1669 (2019).
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).
Alexander, J. M. et al. Live-cell imaging reveals enhancer-dependent Sox2 transcription in the absence of enhancer proximity. eLife 8, e41769 (2019).
Benabdallah, N. S. et al. Decreased enhancer–promoter proximity accompanying enhancer activation. Mol. Cell 76, 473–484 (2019).
Bialek, W., Gregor, T. & Tkačik, G. Action at a distance in transcriptional regulation. Preprint at https://arXiv.org/abs/1912.08579 (2019).
Heist, T., Fukaya, T. & Levine, M. Large distances separate coregulated genes in living Drosophila embryos. Proc. Natl Acad. Sci. USA 116, 15062–15067 (2019).
Richter, W. F., Nayak, S., Iwasa, J. & Taatjes, D. J. The mediator complex as a master regulator of transcription by RNA polymerase II. Nat. Rev. Mol. Cell Biol. 23, 732–749 (2022).
Osman, S. & Cramer, P. Structural biology of RNA polymerase II transcription: 20 years on. Annu. Rev. Cell Dev. Biol. 36, 1–34 (2020).
Soutourina, J. Transcription regulation by the mediator complex. Nat. Rev. Mol. Cell Biol. 19, 262–274 (2018).
Allen, B. L. & Taatjes, D. J. The Mediator complex: a central integrator of transcription. Nat. Rev. Mol. Cell Biol. 16, 155–166 (2015).
Abdella, R. et al. Structure of the human Mediator-bound transcription preinitiation complex. Science 372, 52–56 (2021).
Chen, X. et al. Structures of the human Mediator and Mediator-bound preinitiation complex. Science 372, eabg0635 (2021).
Rengachari, S., Schilbach, S., Aibara, S., Dienemann, C. & Cramer, P. Structure of the human Mediator–RNA polymerase II pre-initiation complex. Nature 594, 129–133 (2021).
Chen, X. et al. Structural insights into preinitiation complex assembly on core promoters. Science 372, eaba8490 (2021).
Panne, D., Maniatis, T. & Harrison, S. C. An atomic model of enhanceosome structure in the vicinity of DNA. Cell 129, 1111 (2007).
Brandão, H. B., Gabriele, M. & Hansen, A. S. Tracking and interpreting long-range chromatin interactions with super-resolution live-cell imaging. Curr. Opin. Cell Biol. 70, 18–26 (2021).
Bellomy, G. R. & Record, M. T. Jr Stable DNA loops in vivo and in vitro: roles in gene regulation at a distance and in biophysical characterization of DNA. Prog. Nucl. Acids Res. Mol. Biol. 39, 81–128 (1990).
Krämer, H., Amouyal, M., Nordheim, A. & Müller-Hill, B. DNA supercoiling changes the spacing requirement of two lac operators for DNA loop formation with lac repressor. EMBO J. 7, 547–556 (1988).
Knight, J. D., Li, R. & Botchan, M. The activation domain of the bovine papillomavirus E2 protein mediates association of DNA-bound dimers to form DNA loops. Proc. Natl Acad. Sci. USA 88, 3204–3208 (1991).
Ptashne, M. & Gann, A. Transcriptional activation by recruitment. Nature 386, 569–577 (1997).
Kyrchanova, O. & Georgiev, P. Mechanisms of enhancer–promoter interactions in higher eukaryotes. Int. J. Mol. Sci. 22, 671 (2021).
Vazquez, J., Muller, M., Pirrotta, V. & Sedat, J. W. The Mcp element mediates stable long-range chromosome–chromosome interactions in Drosophila. Mol. Biol. Cell 17, 2158–2165 (2006).
Merika, M., Williams, A. J., Chen, G., Collins, T. & Thanos, D. Recruitment of CBP/p300 by the IFNβ enhanceosome is required for synergistic activation of transcription. Mol. Cell 1, 277–287 (1998).
Petrenko, N., Jin, Y., Wong, K. H. & Struhl, K. Mediator undergoes a compositional change during transcriptional activation. Mol. Cell 64, 443–454 (2016).
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 (2019).
Du, M. et al. Direct observation of a condensate effect on super-enhancer controlled gene bursting. Cell 187, 1–14 (2024).
Lambert, É., Puwakdandawa, K., Tao, Y. F. & Robert, F. From structure to molecular condensates: emerging mechanisms for mediator function. FEBS J. 90, 286–309 (2023).
Shrinivas, K. et al. Enhancer features that drive formation of transcriptional condensates. Mol. Cell 75, 549–561 (2019).
Li, J. et al. Single-molecule nanoscopy elucidates RNA polymerase II transcription at single genes in live cells. Cell 178, 491–506 (2019).
Boija, A. et al. Transcription factors activate genes through the phase-separation capacity of their activation domains. Cell 175, 1842–1855 (2018).
Lu, H. et al. Phase-separation mechanism for C-terminal hyperphosphorylation of RNA polymerase II. Nature 558, 318–323 (2018).
Cho, W.-K. et al. Mediator and RNA polymerase II clusters associate in transcription-dependent condensates. Science 361, 412–415 (2018).
Sabari, B. R. et al. Coactivator condensation at super-enhancers links phase separation and gene control. Science 361, eaar3958 (2018).
Hu, Z. & Tee, W.-W. Enhancers and chromatin structures: regulatory hubs in gene expression and diseases. Biosci. Rep. 37, BSR20160183 (2017).
Chong, S. et al. Imaging dynamic and selective low-complexity domain interactions that control gene transcription. Science 361, eaar2555 (2018).
Wang, X., Cairns, M. J. & Yan, J. Super-enhancers in transcriptional regulation and genome organization. Nucleic Acids Res. 47, 11481–11496 (2019).
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).
Monfils, K. & Barakat, T. S. Models behind the mystery of establishing enhancer–promoter interactions. Eur. J. Cell Biol. 100, 151170 (2021).
Kent, S. et al. Phase-separated transcriptional condensates accelerate target-search process revealed by live-cell single-molecule imaging. Cell Rep. 33, 108248 (2020).
Gabriele, M. et al. Dynamics of CTCF- and cohesin-mediated chromatin looping revealed by live-cell imaging. Science 376, 496–501 (2022).
Mach, P. et al. Cohesin and CTCF control the dynamics of chromosome folding. Nat. Genet. 54, 1907–1918 (2022).
Horikoshi, M., Hai, T., Lin, Y.-S., Green, M. R. & Roeder, R. G. Transcription factor ATF interacts with the TATA factor to facilitate establishment of a preinitiation complex. Cell 54, 1033–1042 (1988).
Schaffner, W. A hit-and-run mechanism for transcriptional activation? Nature 336, 427–428 (1988).
Pownall, M. E. et al. Chromatin expansion microscopy reveals nanoscale organization of transcription and chromatin. Science 381, 92–100 (2023).
Lammers, N. C., Kim, Y. J., Zhao, J. & Garcia, H. G. A matter of time: using dynamics and theory to uncover mechanisms of transcriptional bursting. Curr. Opin. Cell Biol. 67, 147–157 (2020).
Popp, A. P., Hettich, J. & Gebhardt, J. C. M. Altering transcription factor binding reveals comprehensive transcriptional kinetics of a basic gene. Nucleic Acids Res. 49, 6249–6266 (2021).
Stavreva, D. A. et al. Transcriptional bursting and co-bursting regulation by steroid hormone release pattern and transcription factor mobility. Mol. Cell 75, 1161–1177 (2019).
Fritzsch, C. et al. Estrogen-dependent control and cell-to-cell variability of transcriptional bursting. Mol. Syst. Biol. 14, e7678 (2018).
Tantale, K. et al. A single-molecule view of transcription reveals convoys of RNA polymerases and multi-scale bursting. Nat. Commun. 7, 12248 (2016).
Teves, S. S. et al. A dynamic mode of mitotic bookmarking by transcription factors. eLife 5, e22280 (2016).
Larson, D. R. et al. Direct observation of frequency modulated transcription in single cells using light activation. eLife 2, e00750 (2013).
Mazza, D., Abernathy, A., Golob, N., Morisaki, T. & McNally, J. G. A benchmark for chromatin binding measurements in live cells. Nucleic Acids Res. 40, e119 (2012).
Suter, D. M. et al. Mammalian genes are transcribed with widely different bursting kinetics. Science 332, 472–474 (2011).
McNally, J. G., Muller, W. G., Walker, D., Wolford, R. & Hager, G. L. The glucocorticoid receptor: rapid exchange with regulatory sites in living cells. Science 287, 1262–1265 (2000).
Zhang, Q., Shi, H. & Zhang, Z. A dynamic kissing model for enhancer–promoter communication on the surface of transcriptional condensate. Preprint at bioRxiv https://doi.org/10.1101/2022.03.03.482814 (2022).
Baek, I., Friedman, L. J., Gelles, J. & Buratowski, S. Single-molecule studies reveal branched pathways for activator-dependent assembly of RNA polymerase II pre-initiation complexes. Mol. Cell 81, 3576–3588 (2021).
Thomas, H. F. et al. Temporal dissection of an enhancer cluster reveals distinct temporal and functional contributions of individual elements. Mol. Cell 81, 969–982 (2021).
Hou, C., Zhao, H., Tanimoto, K. & Dean, A. CTCF-dependent enhancer-blocking by alternative chromatin loop formation. Proc. Natl Acad. Sci. USA 105, 20398–20403 (2008).
Andersson, R. & Sandelin, A. Determinants of enhancer and promoter activities of regulatory elements. Nat. Rev. Genet. 21, 71–87 (2020).
Buckley, M. S. & Lis, J. T. Imaging RNA polymerase II transcription sites in living cells. Curr. Opin. Genet. Dev. 25, 126–130 (2014).
Kubo, N. et al. Promoter-proximal CTCF binding promotes distal enhancer-dependent gene activation. Nat. Struct. Mol. Biol. 28, 152–161 (2021).
Gibbons, M. D. et al. Enhancer-mediated formation of nuclear transcription initiation domains. Int. J. Mol. Sci. 23, 9290 (2022).
Reiter, F., Wienerroither, S. & Stark, A. Combinatorial function of transcription factors and cofactors. Curr. Opin. Genet. Dev. 43, 73–81 (2017).
Shlyueva, D., Stampfel, G. & Stark, A. Transcriptional enhancers: from properties to genome-wide predictions. Nat. Rev. Genet. 15, 272–286 (2014).
Narita, T. et al. Enhancers are activated by p300/CBP activity-dependent PIC assembly, RNAPII recruitment, and pause release. Mol. Cell 81, 2166–2182 (2021).
Hsu, E., Zemke, N. R. & Berk, A. J. Promoter-specific changes in initiation, elongation, and homeostasis of histone H3 acetylation during CBP/p300 inhibition. eLife 10, e63512 (2021).
Core, L. & Adelman, K. Promoter-proximal pausing of RNA polymerase II: a nexus of gene regulation. Genes Dev. 33, 960–982 (2019).
Chen, F. X., Smith, E. R. & Shilatifard, A. Born to run: control of transcription elongation by RNA polymerase II. Nat. Rev. Mol. Cell Biol. 19, 464–478 (2018).
Mir, M. et al. Dynamic multifactor hubs interact transiently with sites of active transcription in Drosophila embryos. eLife 7, e40497 (2018).
Hansen, A. S., Amitai, A., Cattoglio, C., Tjian, R. & Darzacq, X. Guided nuclear exploration increases CTCF target search efficiency. Nat. Chem. Biol. 16, 257–266 (2020).
Trojanowski, J. et al. Transcription activation is enhanced by multivalent interactions independent of phase separation. Mol. Cell 82, 1878–1893 (2022).
Chong, S. et al. Tuning levels of low-complexity domain interactions to modulate endogenous oncogenic transcription. Mol. Cell 82, 2084–2097 (2022).
Panigrahi, A. & O’Malley, B. W. Mechanisms of enhancer action: the known and the unknown. Genome Biol. 22, 1–30 (2021).
Malik, S. & Roeder, R. G. Mediator: a drawbridge across the enhancer–promoter divide. Mol. Cell 64, 433–434 (2016).
Kim, Y.-J., Björklund, S., Li, Y., Sayre, M. H. & Kornberg, R. D. A multiprotein mediator of transcriptional activation and its interaction with the c-terminal repeat domain of RNA polymerase II. Cell 77, 599–608 (1994).
Chen, Q. et al. Enhancer RNAs in transcriptional regulation: recent insights. Front. Cell Dev. Biol. 11, 1205540 (2023).
Zhao, Y. et al. Activation of P-TEFb by androgen receptor-regulated enhancer RNAs in castration-resistant prostate cancer. Cell Rep. 15, 599–610 (2016).
Xiao, J. Y., Hafner, A. & Boettiger, A. N. How subtle changes in 3D structure can create large changes in transcription. eLife 10, e64320 (2021).
Tsujimura, T. et al. Controlling gene activation by enhancers through a drug-inducible topological insulator. eLife 9, e47980 (2020).
Wu, H., Zhang, J., Tan, L. & Xie, X. S. Extruding transcription elongation loops observed in high-resolution single-cell 3D genomes. Preprint at bioRxiv https://doi.org/10.1101/2023.02.18.529096 (2023).
Danino, Y. M., Even, D., Ideses, D. & Juven-Gershon, T. The core promoter: at the heart of gene expression. Biochim. Biophys. Acta Gene Regul. Mech. 1849, 1116–1131 (2015).
Wang, Z. et al. Prediction of histone post-translational modification patterns based on nascent transcription data. Nat. Genet. 54, 295–305 (2022).
Zheng, Y., Thomas, P. M. & Kelleher, N. L. Measurement of acetylation turnover at distinct lysines in human histones identifies long-lived acetylation sites. Nat. Commun. 4, 2203 (2013).
Claringbould, A. & Zaugg, J. B. Enhancers in disease: molecular basis and emerging treatment strategies. Trends Mol. Med. 27, 1060–1073 (2021).
Jia, Q., Chen, S., Tan, Y., Li, Y. & Tang, F. Oncogenic super-enhancer formation in tumorigenesis and its molecular mechanisms. Exp. Mol. Med. 52, 713–723 (2020).
Beroukhim, R., Zhang, X. & Meyerson, M. Copy number alterations unmasked as enhancer hijackers. Nat. Genet. 49, 5–6 (2017).
Herz, H.-M. Enhancer deregulation in cancer and other diseases. BioEssays 38, 1003–1015 (2016).
Northcott, P. A. et al. Enhancer hijacking activates GFI1 family oncogenes in medulloblastoma. Nature 511, 428–434 (2014).
Flavahan, W. A. et al. Insulator dysfunction and oncogene activation in IDH mutant gliomas. Nature 529, 110–114 (2016).
Flavahan, W. A. et al. Altered chromosomal topology drives oncogenic programs in SDH-deficient GISTs. Nature 575, 229–233 (2019).
Klemm, S. L., Shipony, Z. & Greenleaf, W. J. Chromatin accessibility and the regulatory epigenome. Nat. Rev. Genet. 20, 207–220 (2019).
Bell, O., Tiwari, V. K., Thomä, N. H. & Schübeler, D. Determinants and dynamics of genome accessibility. Nat. Rev. Genet. 12, 554–564 (2011).
Starks, R. R., Biswas, A., Jain, A. & Tuteja, G. Combined analysis of dissimilar promoter accessibility and gene expression profiles identifies tissue-specific genes and actively repressed networks. Epigenetics Chromatin 12, 1–16 (2019).
Szyf, M., Weaver, I. & Meaney, M. Maternal care, the epigenome and phenotypic differences in behavior. Reprod. Toxicol. 24, 9–19 (2007).
Esteller, M. Epigenetic gene silencing in cancer: the DNA hypermethylome. Hum. Mol. Genet. 16, R50–R59 (2007).
Issa, J.-P. CpG island methylator phenotype in cancer. Nat. Rev. Cancer 4, 988–993 (2004).
Tycko, B. et al. Epigenetic gene silencing in cancer. J. Clin. Invest. 105, 401–407 (2000).
Lee, K. et al. Integrated analysis of tissue-specific promoter methylation and gene expression profile in complex diseases. Int. J. Mol. Sci. 21, 5056 (2020).
Schilling, E. & Rehli, M. Global, comparative analysis of tissue-specific promoter CpG methylation. Genomics 90, 314–323 (2007).
Perissi, V., Jepsen, K., Glass, C. K. & Rosenfeld, M. G. Deconstructing repression: evolving models of co-repressor action. Nat. Rev. Genet. 11, 109–123 (2010).
Payankaulam, S., Li, L. M. & Arnosti, D. N. Transcriptional repression: conserved and evolved features. Curr. Biol. 20, R764–R771 (2010).
Blackledge, N. P. & Klose, R. J. The molecular principles of gene regulation by Polycomb repressive complexes. Nat. Rev. Mol. Cell Biol. 22, 815–833 (2021).
Cheutin, T. & Cavalli, G. Polycomb silencing: from linear chromatin domains to 3D chromosome folding. Curr. Opin. Genet. Dev. 25, 30–37 (2014).
Zhang, Y., See, Y. X., Tergaonkar, V. & Fullwood, M. J. Long-distance repression by human silencers: chromatin interactions and phase separation in silencers. Cells 11, 1560 (2022).
Cornejo-Páramo, P., Roper, K., Degnan, S. M., Degnan, B. M. & Wong, E. S. Distal regulation, silencers, and a shared combinatorial syntax are hallmarks of animal embryogenesis. Genome Res. 32, 474–487 (2022).
Courey, A. J. & Jia, S. Transcriptional repression: the long and the short of it. Genes Dev. 15, 2786–2796 (2001).
Burke, L. J. & Baniahmad, A. Co-repressors 2000. FASEB J. 14, 1876–1888 (2000).
Altincicek, B. et al. Interaction of the corepressor alien with DAX-1 is abrogated by mutations of DAX-1 involved in adrenal hypoplasia congenita. J. Biol. Chem. 275, 7662–7667 (2000).
Amir, R. E. et al. Rett syndrome is caused by mutations in X-linked MECP2, encoding methyl-CpG-binding protein 2. Nat. Genet. 23, 185–188 (1999).
Muscatelli, F. et al. Mutations in the DAX-1 gene give rise to both X-linked adrenal hypoplasia congenita and hypogonadotropic hypogonadism. Nature 372, 672–676 (1994).
Jacobs, J., Pagani, M., Wenzl, C. & Stark, A. Widespread regulatory specificities between transcriptional co-repressors and enhancers in Drosophila. Science 381, 198–204 (2023).
Bozek, M. & Gompel, N. Developmental transcriptional enhancers: a subtle interplay between accessibility and activity: considering quantitative accessibility changes between different regulatory states of an enhancer deconvolutes the complex relationship between accessibility and activity. BioEssays 42, 1900188 (2020).
Li, X. & Noll, M. Compatibility between enhancers and promoters determines the transcriptional specificity of gooseberry and gooseberry neuro in the Drosophila embryo. EMBO J. 13, 400–406 (1994).
Juven-Gershon, T., Hsu, J.-Y. & Kadonaga, J. T. Caudal, a key developmental regulator, is a DPE-specific transcriptional factor. Genes Dev. 22, 2823–2830 (2008).
Butler, J. E. & Kadonaga, J. T. Enhancer–promoter specificity mediated by DEP or TATA core promoter motifs. Genes Dev. 15, 2515–2519 (2001).
Ohtsuki, S., Levine, M. & Cai, H. N. Different core promoters possess distinct regulatory activities in the Drosophila embryo. Genes Dev. 12, 547–556 (1998).
Kwon, D. et al. Enhancer–promoter communication at the Drosophila engrailed locus. Development 136, 3067–3075 (2009).
Akbari, O. S. et al. A novel promoter-tethering element regulates enhancer-driven gene expression at the bithorax complex in the Drosophila embryo. Development 135, 123–131 (2008).
Shir-Shapira, H. et al. Identification of evolutionarily conserved downstream core promoter elements required for the transcriptional regulation of Fushi tarazu target genes. PLoS ONE 14, e0215695 (2019).
Juven-Gershon, T. & Kadonaga, J. T. Regulation of gene expression via the core promoter and the basal transcriptional machinery. Dev. Biol. 339, 225–229 (2010).
Natsume-Kitatani, Y. & Mamitsuka, H. Classification of promoters based on the combination of core promoter elements exhibits different histone modification patterns. PLoS ONE 11, e0151917 (2016).
Cermakova, K. & Hodges, H. C. Interaction modules that impart specificity to disordered protein. Trends Biochem. Sci. 48, 477–490 (2023).
Chong, S. & Mir, M. Towards decoding the sequence-based grammar governing the functions of intrinsically disordered protein regions. J. Mol. Biol. 433, 166724 (2021).
Brodsky, S. et al. Intrinsically disordered regions direct transcription factor in vivo binding specificity. Mol. Cell 79, 459–471 (2020).
Bergman, D. T. et al. Compatibility rules of human enhancer and promoter sequences. Nature 607, 176–184 (2022).
Martinez-Ara, M., Comoglio, F., van Arensbergen, J. & van Steensel, B. Systematic analysis of intrinsic enhancer–promoter compatibility in the mouse genome. Mol. Cell 82, 2519–2531 (2022).
Wang, H.-L. V. & Corces, V. G. The cupid shuffle: do enhancers prefer specific promoters? Mol. Cell 82, 2357–2359 (2022).
Sahu, B. et al. Sequence determinants of human gene regulatory elements. Nat. Genet. 54, 283–294 (2022).
Nora, E. P. et al. Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 485, 381–385 (2012).
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).
Rao, S. S. et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159, 1665–1680 (2014).
Golfier, S., Quail, T., Kimura, H. & Brugués, J. Cohesin and condensin extrude DNA loops in a cell cycle-dependent manner. eLife 9, e53885 (2020).
Davidson, I. F. et al. DNA loop extrusion by human cohesin. Science 366, 1338–1345 (2019).
Kim, Y., Shi, Z., Zhang, H., Finkelstein, I. J. & Yu, H. Human cohesin compacts DNA by loop extrusion. Science 366, 1345–1349 (2019).
Ganji, M. et al. Real-time imaging of DNA loop extrusion by condensin. Science 360, 102–105 (2018).
Fudenberg, G. et al. Formation of chromosomal domains by loop extrusion. Cell Rep. 15, 2038–2049 (2016).
Rao, S. S. et al. Cohesin loss eliminates all loop domains. Cell 171, 305–320 (2017).
Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944 (2017).
Schwarzer, W. et al. Two independent modes of chromatin organization revealed by cohesin removal. Nature 551, 51–56 (2017).
Gassler, J. et al. A mechanism of cohesin-dependent loop extrusion organizes zygotic genome architecture. EMBO J. 36, 3600–3618 (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).
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).
de Wit, E. et al. CTCF binding polarity determines chromatin looping. Mol. Cell 60, 676–684 (2015).
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).
Symmons, O. et al. Functional and topological characteristics of mammalian regulatory domains. Genome Res. 24, 390–400 (2014).
Arrastia, M. V. et al. Single-cell measurement of higher-order 3D genome organization with scSPRITE. Nat. Biotechnol. 40, 64–73 (2022).
Wutz, G. et al. ESCO1 and CTCF enable formation of long chromatin loops by protecting cohesinSTAG1 from WAPL. eLife 9, e52091 (2020).
Vian, L. et al. The energetics and physiological impact of cohesin extrusion. Cell 173, 1165–1178 (2018).
Dekker, J. & Mirny, L. The 3D genome as moderator of chromosomal communication. Cell 164, 1110–1121 (2016).
Doyle, B., Fudenberg, G., Imakaev, M. & Mirny, L. A. Chromatin loops as allosteric modulators of enhancer–promoter interactions. PLoS Comput. Biol. 10, e1003867 (2014).
Oh, S. et al. Enhancer release and retargeting activates disease-susceptibility genes. Nature 595, 735–740 (2021).
Ealo, T. et al. Synergistic insulation of regulatory domains by developmental genes and clusters of CTCF sites. Preprint at bioRxiv https://doi.org/10.1101/2023.12.15.571760 (2023).
Lupiez, D. G. et al. Disruptions of topological chromatin domains cause pathogenic rewiring of gene–enhancer interactions. Cell 161, 1012–1025 (2015).
Paliou, C. et al. Preformed chromatin topology assists transcriptional robustness of Shh during limb development. Proc. Natl Acad. Sci. USA 116, 12390–12399 (2019).
Schuijers, J. et al. Transcriptional dysregulation of MYC reveals common enhancer-docking mechanism. Cell Rep. 23, 349–360 (2018).
Geyer, P. K. & Corces, V. G. DNA position-specific repression of transcription by a Drosophila zinc finger protein. Genes Dev. 6, 1865–1873 (1992).
Kellum, R. & Schedl, P. A position-effect assay for boundaries of higher order chromosomal domains. Cell 64, 941–950 (1991).
Udvardy, A., Maine, E. & Schedl, P. The 87A7 chromomere: identification of novel chromatin structures flanking the heat shock locus that may define the boundaries of higher order domains. J. Mol. Biol. 185, 341–358 (1985).
Kyrchanova, O., Sokolov, V. & Georgiev, P. Mechanisms of interaction between enhancers and promoters in three Drosophila model systems. Int. J. Mol. Sci. 24, 2855 (2023).
Batut, P. J. et al. Genome organization controls transcriptional dynamics during development. Science 375, 566–570 (2022).
Deng, H., Jin, G. & Lim, B. Unveiling dynamic enhancer–promoter interactions in Drosophila melanogaster. Biochem. Soc. Trans. 50, 1633–1642 (2022).
Kyrchanova, O. & Georgiev, P. Chromatin insulators and long-distance interactions in Drosophila. FEBS Lett. 588, 8–14 (2014).
da Costa-Nunes, J. A. & Noordermeer, D. Tads: dynamic structures to create stable regulatory functions. Curr. Opin. Struct. Biol. 81, 102622 (2023).
Akgol Oksuz, B. et al. Systematic evaluation of chromosome conformation capture assays. Nat. Methods 18, 1046–1055 (2021).
Fudenberg, G. & Imakaev, M. FISH-ing for captured contacts: towards reconciling FISH and 3C. Nat. Methods 14, 673–678 (2017).
Krietenstein, N. et al. Ultrastructural details of mammalian chromosome architecture. Mol. Cell 78, 554–565 (2020).
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).
Robles-Rebollo, I. et al. Cohesin couples transcriptional bursting probabilities of inducible enhancers and promoters. Nat. Commun. 13, 4342 (2022).
Cuartero, S. et al. Control of inducible gene expression links cohesin to hematopoietic progenitor self-renewal and differentiation. Nat. Immunol. 19, 932–941 (2018).
Kane, L. et al. Cohesin is required for long-range enhancer action at the Shh locus. Nat. Struct. Mol. Biol. 29, 891–897 (2022).
Calderon, L. et al. Cohesin-dependence of neuronal gene expression relates to chromatin loop length. eLife 11, e76539 (2022).
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).
Marshall, W. et al. Interphase chromosomes undergo constrained diffusional motion in living cells. Curr. Biol. 7, 930–939 (1997).
Keizer, V. I. et al. Live-cell micromanipulation of a genomic locus reveals interphase chromatin mechanics. Science 377, 489–495 (2022).
Bénichou, O., Guérin, T. & Voituriez, R. Mean first-passage times in confined media: from Markovian to non-Markovian processes. J. Phys. A Math. Theor. 48, 163001 (2015).
Yang, J. H., Brandão, H. B. & Hansen, A. S. DNA double-strand break end synapsis by DNA loop extrusion. Nat. Commun. 14, 1913 (2023).
Cattoglio, C. et al. Determining cellular CTCF and cohesin abundances to constrain 3D genome models. eLife 8, e40164 (2019).
Holzmann, J. et al. Absolute quantification of cohesin, CTCF and their regulators in human cells. eLife 8, e46269 (2019).
Hamamoto, K. & Fukaya, T. Molecular architecture of enhancer–promoter interaction. Curr. Opin. Cell Biol. 74, 62–70 (2022).
Greenwald, W. W. et al. Subtle changes in chromatin loop contact propensity are associated with differential gene regulation and expression. Nat. Commun. 10, 1054 (2019).
Whalen, S., Truty, R. M. & Pollard, K. S. Enhancer–promoter interactions are encoded by complex genomic signatures on looping chromatin. Nat. Genet. 48, 488–496 (2016).
Hyle, J. et al. Acute depletion of CTCF directly affects MYC regulation through loss of enhancer–promoter looping. Nucleic Acids Res. 47, 6699–6713 (2019).
Cerda-Smith, C. et al. Integrative PTEN enhancer discovery reveals a new model of enhancer organization. Preprint at bioRxiv https://doi.org/10.1101/2023.09.20.558459 (2023).
Zhang, X. et al. Identification of focally amplified lineage-specific super-enhancers in human epithelial cancers. Nat. Genet. 48, 176–182 (2016).
Korch, C. et al. DNA profiling analysis of endometrial and ovarian cell lines reveals misidentification, redundancy and contamination. Gynecol. Oncol. 127, 241–248 (2012).
He, B., Chen, C., Teng, L. & Tan, K. Global view of enhancer–promoter interactome in human cells. Proc. Natl Acad. Sci. USA 111, E2191–E2199 (2014).
Grosse-Holz, S., Coulon, A. & Mirny, L. Scale-free models of chromosome structure, dynamics, and mechanics. Preprint at bioRxiv https://doi.org/10.1101/2023.04.14.536939 (2023).
Giorgetti, L. & Heard, E. Closing the loop: 3C versus DNA FISH. Genome Biol. 17, 1–9 (2016).
Kempfer, R. & Pombo, A. Methods for mapping 3D chromosome architecture. Nat. Rev. Genet. 21, 207–226 (2020).
Cremer, T. & Cremer, C. Chromosome territories, nuclear architecture and gene regulation in mammalian cells. Nat. Rev. Genet. 2, 292–301 (2001).
Bridger, J. M. & Volpi, E. V. Fluorescence in situ Hybridization (FISH): Protocols and Applications (Humana Press, 2010).
Speicher, M. R., Ballard, S. G. & Ward, D. C. Karyotyping human chromosomes by combinatorial multi-fluor FISH. Nat. Genet. 12, 368–375 (1996).
Gall, J. G. & Pardue, M. L. Formation and detection of RNA–DNA hybrid molecules in cytological preparations. Proc. Natl Acad. Sci. USA 63, 378–383 (1969).
Liu, M. et al. Multiplexed imaging of nucleome architectures in single cells of mammalian tissue. Nat. Commun. 11, 2907 (2020).
Sawh, A. N. et al. Lamina-dependent stretching and unconventional chromosome compartments in early C. elegans embryos. Mol. Cell 78, 96–111 (2020).
Gizzi, A. M. C. et al. Microscopy-based chromosome conformation capture enables simultaneous visualization of genome organization and transcription in intact organisms. Mol. Cell 74, 212–222 (2019).
Bintu, B. et al. Super-resolution chromatin tracing reveals domains and cooperative interactions in single cells. Science 362, eaau1783 (2018).
Nir, G. et al. Walking along chromosomes with super-resolution imaging, contact maps, and integrative modeling. PLoS Genet. 14, e1007872 (2018).
Wang, S. et al. Spatial organization of chromatin domains and compartments in single chromosomes. Science 353, 598–602 (2016).
Beliveau, B. J. et al. Versatile design and synthesis platform for visualizing genomes with Oligopaint FISH probes. Proc. Natl Acad. Sci. USA 109, 21301–21306 (2012).
Irgen-Gioro, S., Yoshida, S., Walling, V. & Chong, S. Fixation can change the appearance of phase separation in living cells. eLife 11, e79903 (2022).
Beckwith, K. et al. Nanoscale 3D DNA tracing in single human cells visualizes loop extrusion directly in situ. Preprint at bioRxiv https://doi.org/10.1101/2021.04.12.439407 (2021).
Brown, J. M., De Ornellas, S., Parisi, E., Schermelleh, L. & Buckle, V. J. RASER-FISH: non-denaturing fluorescence in situ hybridization for preservation of three-dimensional interphase chromatin structure. Nat. Protoc. 17, 1306–1331 (2022).
Lakadamyali, M. & Cosma, M. P. Visualizing the genome in high resolution challenges our textbook understanding. Nat. Methods 17, 371–379 (2020).
Khanna, N., Zhang, Y., Lucas, J. S., Dudko, O. K. & Murre, C. Chromosome dynamics near the sol–gel phase transition dictate the timing of remote genomic interactions. Nat. Commun. 10, 2771 (2019).
Germier, T. et al. Real-time imaging of a single gene reveals transcription-initiated local confinement. Biophys. J. 113, 1383–1394 (2017).
Robinett, C. C. et al. In vivo localization of DNA sequences and visualization of large-scale chromatin organization using lac operator/repressor recognition. J. Cell Biol. 135, 1685–1700 (1996).
Platania, A. et al. Competition between transcription and loop extrusion modulates promoter and enhancer dynamics. Preprint at bioRxiv https://doi.org/10.1101/2023.04.25.538222 (2023).
Bohrer, C. H. & Larson, D. R. Synthetic analysis of chromatin tracing and live-cell imaging indicates pervasive spatial coupling between genes. eLife 12, e81861 (2023).
Acuña, L. I. G., Flyamer, I., Boyle, S., Friman, E. & Bickmore, W. A. Transcription decouples estrogen-dependent changes in enhancer–promoter contact frequencies and spatial proximity. Preprint at bioRxiv https://doi.org/10.1101/2023.03.29.534720 (2023).
Barinov, L., Ryabichko, S., Bialek, W. & Gregor, T. Transcription-dependent spatial organization of a gene locus. Preprint at https://arXiv.org/abs/2012.15819 (2020).
Thompson, R. E., Larson, D. R. & Webb, W. W. Precise nanometer localization analysis for individual fluorescent probes. Biophys. J. 82, 2775–2783 (2002).
Imakaev, M. et al. Iterative correction of Hi-C data reveals hallmarks of chromosome organization. Nat. Methods 9, 999–1003 (2012).
Acknowledgements
The authors thank L. Mirny, S. Grosse-Holz, H. Pinholt, M. Mir, C. Hong, M. Gabriele, M. Mazzocca and the rest of the Hansen laboratory for insightful discussions and comments on the manuscript. J.H.Y. was supported by the MathWorks Engineering Fellowship and a graduate fellowship from the Ludwig Center at MIT’s Koch Institute for Integrative Cancer Research. This work was supported by the National Institutes of Health (grant numbers R00GM130896, DP2GM140938, R33CA257878, UM1HG011536), the National Science Foundation (grant 2036037) and a Pew-Stewart Cancer Research Scholar grant.
Author information
Authors and Affiliations
Contributions
J.H.Y. researched data for the article. Both authors contributed substantially to discussion of the content, wrote the article and reviewed and edited the manuscript before submission.
Corresponding author
Ethics declarations
Competing interests
The authors declare no competing interests.
Peer review
Peer review information
Nature Reviews Molecular Cell Biology thanks Thomas Milne, Marieke Oudelaar with Abrar Aljahani, and Ranjith Padinhateeri 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.
Supplementary information
Glossary
- Architectural proteins
-
Proteins that regulate 3D chromatin structure by forming chromatin loops and domains, which can regulate interactions between enhancers and promoters.
- Biochemical compatibility
-
The intrinsic ability for an enhancer to activate transcription at some promoters but not others, which may be determined by the binding profile of transcription factors and co-activators.
- Chromatic aberrations
-
Owing to the refractive index varying with the wavelength of light, a perfectly colocalizing E–P pair (true distance of 0 nm) may be measured as being far apart. Very accurate correction of chromatic aberrations is required for precise measurements of E–P distances.
- Clustering
-
A cluster corresponds to higher-than-expected local density of molecules. The term cluster is agnostic to the mechanism of cluster formation and clusters are often defined using spatial statistics.
- Condensates
-
Refers to formation of membraneless compartments of high local concentration of factors through liquid–liquid phase separation.
- Co-repressors
-
Enzymatic complexes recruited to DNA directly or indirectly by transcription factors to establish and maintain repression of transcription.
- Enhanceosome
-
A protein complex that assembles on an enhancer to regulate the transcription of the cognate promoter.
- Enhancer RNAs
-
Non-coding RNAs transcribed from enhancers, which may have gene regulatory functions.
- E–P interaction radius
-
The maximum 3D distance between an enhancer and a promoter that enables them to functionally interact.
- Facilitator
-
Refers to DNA–protein complexes such as DNA-bound CTCF, which can increase the interaction probability between promoters and regulatory elements such as enhancers and silencers.
- First-passage time
-
The time taken for a stochastic process to reach a specific state for the first time, for example, the time taken for an enhancer to find and interact with a promoter.
- Hub
-
Discrete nuclear domains of high transcription protein concentration, which serve as a focal point of activity; ‘hub formation’ is often used to indicate a formation mechanism that is distinct from phase separation.
- Insulators
-
DNA elements bound by specific protein complexes, which may reduce gene expression when placed between an enhancer and a promoter, presumably by reducing the probability of their interaction.
- Intrinsically disordered regions
-
Protein segments lacking well-defined 3D structure in physiological conditions, which form dynamic ensembles of conformations and may engage in multivalent interactions.
- Mean squared displacement
-
The average of the squared displacement of a particle or locus with respect to a reference position, usually calculated over a range of time intervals.
- Rouse dynamics
-
The movement and behaviour of polymers in a bead–spring model, in which monomers are connected by Hookean springs and a monomer only interacts with its nearest neighbours.
- Silencers
-
Regulatory DNA elements that reduce transcription at cognate promoters, including from far away in the genome.
- Super-enhancers
-
Genomic regions consisting of multiple enhancers that can drive high level of transcription at cognate promoters; originally defined based on Mediator enrichment.
- Transcription memory
-
The phenomenon in which the influence of a stimulus persists beyond the initial exposure to the stimulus, including promoter memory of past E–P interactions.
Rights and permissions
Springer Nature or its licensor (e.g. a society or other partner) 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.
About this article
Cite this article
Yang, J.H., Hansen, A.S. Enhancer selectivity in space and time: from enhancer–promoter interactions to promoter activation. Nat Rev Mol Cell Biol (2024). https://doi.org/10.1038/s41580-024-00710-6
Accepted:
Published:
DOI: https://doi.org/10.1038/s41580-024-00710-6
