Abstract
Spatiotemporal gene expression programmes are orchestrated by transcriptional enhancers, which are key regulatory DNA elements that engage in physical contacts with their target-gene promoters, often bridging considerable genomic distances. Recent progress in genomics, genome editing and microscopy methodologies have enabled the genome-wide mapping of enhancer–promoter contacts and their functional dissection. In this Review, we discuss novel concepts on how enhancer–promoter interactions are established and maintained, how the 3D architecture of mammalian genomes both facilitates and constrains enhancer–promoter contacts, and the role they play in gene expression control during normal development and disease.
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References
Plank, J. L. & Dean, A. Enhancer function: mechanistic and genome-wide insights come together. Mol. Cell 55, 5–14 (2014).
Pombo, A. & Dillon, N. Three-dimensional genome architecture: players and mechanisms. Nat. Rev. Mol. Cell Biol. 16, 245–257 (2015).
Long, H. K., Prescott, S. L. & Wysocka, J. Ever-changing landscapes: transcriptional enhancers in development and evolution. Cell 167, 1170–1187 (2016).
Furlong, E. E. M. & Levine, M. Developmental enhancers and chromosome topology. Science 361, 1341–1345 (2018).
Lettice, L. A. et al. A long-range Shh enhancer regulates expression in the developing limb and fin and is associated with preaxial polydactyly. Hum. Mol. Genet. 12, 1725–1735 (2003). This work identified the long-range ZRS enhancer driving the expression of Shh in the developing limb bud and demonstrated that point mutations in this enhancer can cause congenital limb malformations.
Benko, S. et al. Highly conserved non-coding elements on either side of SOX9 associated with Pierre Robin sequence. Nat. Genet. 41, 359–364 (2009).
Bhatia, S. et al. Disruption of autoregulatory feedback by a mutation in a remote, ultraconserved PAX6 enhancer causes aniridia. Am. J. Hum. Genet. 93, 1126–1134 (2013).
Uslu, V. V. et al. Long-range enhancers regulating Myc expression are required for normal facial morphogenesis. Nat. Genet. 46, 753–758 (2014).
Lupianez, D. G. et al. Disruptions of topological chromatin domains cause pathogenic rewiring of gene-enhancer interactions. Cell 161, 1012–1025 (2015). This paper demonstrated that genomic rearrangements that disrupt TAD integrity can result in ectopic enhancer–promoter interactions that cause defects during human limb development.
Franke, M. et al. Formation of new chromatin domains determines pathogenicity of genomic duplications. Nature 538, 265–269 (2016).
Shen, Y. et al. A map of the cis-regulatory sequences in the mouse genome. Nature 488, 116–120 (2012).
ENCODE Project Consortium. An integrated encyclopedia of DNA elements in the human genome. Nature 489, 57–74 (2012).
Amano, T. et al. Chromosomal dynamics at the Shh locus: limb bud-specific differential regulation of competence and active transcription. Dev. Cell 16, 47–57 (2009).
Sanyal, A. et al. The long-range interaction landscape of gene promoters. Nature 489, 109–113 (2012).
Schoenfelder, S. et al. The pluripotent regulatory circuitry connecting promoters to their long-range interacting elements. Genome Res. 25, 582–597 (2015).
Mifsud, B. et al. Mapping long-range promoter contacts in human cells with high-resolution capture Hi-C. Nat. Genet. 47, 598–606 (2015). Schoenfelder et al. ( Genome Res ., 2015) and Mifsud et al. (2015) established promoter-capture Hi-C for genome-wide profiling of promoter interactions at high resolution for all promoters in the mouse (Schoenfelder et al. ( Genome Res ., 2015)) and human (Mifsud et al. (2015)) genomes.
Wilson, N. K. et al. Integrated genome-scale analysis of the transcriptional regulatory landscape in a blood stem/progenitor cell model. Blood 127, e12–e23 (2016).
Javierre, B. M. et al. Lineage-specific genome architecture links enhancers and non-coding disease variants to target gene promoters. Cell 167, 1369–1384 (2016). By mapping long-range promoter interactions in 17 primary human blood cell types, this study linked thousands of non-coding GWAS SNPs to their putative target genes.
Siersbaek, R. et al. Dynamic rewiring of promoter-anchored chromatin loops during adipocyte differentiation. Mol. Cell 66, 420–435 (2017).
Freire-Pritchett, P. et al. Global reorganisation of cis-regulatory units upon lineage commitment of human embryonic stem cells. eLife 6, e21926 (2017).
Rubin, A. J. et al. Lineage-specific dynamic and pre-established enhancer-promoter contacts cooperate in terminal differentiation. Nat. Genet. 49, 1522–1528 (2017).
Novo, C. L. et al. Long-range enhancer interactions are prevalent in mouse embryonic stem cells and are reorganized upon pluripotent state transition. Cell Rep. 22, 2615–2627 (2018).
Schoenfelder, S. et al. Divergent wiring of repressive and active chromatin interactions between mouse embryonic and trophoblast lineages. Nat. Commun. 9, 4189 (2018).
Sanjana, N. E. et al. High-resolution interrogation of functional elements in the noncoding genome. Science 353, 1545–1549 (2016).
Rajagopal, N. et al. High-throughput mapping of regulatory DNA. Nat. Biotechnol. 34, 167–174 (2016).
Fulco, C. P. et al. Systematic mapping of functional enhancer-promoter connections with CRISPR interference. Science 354, 769–773 (2016).
Diao, Y. et al. A new class of temporarily phenotypic enhancers identified by CRISPR/Cas9-mediated genetic screening. Genome Res. 26, 397–405 (2016).
Diao, Y. et al. A tiling-deletion-based genetic screen for cis-regulatory element identification in mammalian cells. Nat. Methods 14, 629–635 (2017).
Klann, T. S. et al. CRISPR-Cas9 epigenome editing enables high-throughput screening for functional regulatory elements in the human genome. Nat. Biotechnol. 35, 561–568 (2017).
Simeonov, D. R. et al. Discovery of stimulation-responsive immune enhancers with CRISPR activation. Nature 549, 111–115 (2017).
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). This study is the first to demonstrate that an induced juxtaposition of an enhancer with its target promoter can induce gene transcription.
Deng, W. et al. Reactivation of developmentally silenced globin genes by forced chromatin looping. Cell 158, 849–860 (2014).
Bartman, C. R., Hsu, S. C., Hsiung, C. C., Raj, A. & Blobel, G. A. Enhancer regulation of transcriptional bursting parameters revealed by forced chromatin looping. Mol. Cell 62, 237–247 (2016).
Haarhuis, J. H. I. et al. The cohesin release factor WAPL restricts chromatin loop extension. Cell 169, 693–707 (2017).
Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944 (2017). Using an acute protein depletion system, this study shows that CTCF is required for looping between CTCF target sites and TAD insulation but dispensable for the segregation of the genome into active and inactive compartments.
Schwarzer, W. et al. Two independent modes of chromatin organization revealed by cohesin removal. Nature 551, 51–56 (2017).
Rao, S. S. P. et al. Cohesin loss eliminates all loop domains. Cell 171, 305–320 (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). Schwarzer et al. (2017), Rao et al. (2017) and Wutz et al. (2017) show that cohesin is required for TADs and loops but not compartments; unexpectedly, cohesin depletion has only mild effects on gene expression genome-wide.
Weintraub, A. S. et al. YY1 Is a structural regulator of enhancer-promoter loops. Cell 171, 1573–1588 (2017).
Spitz, F. Gene regulation at a distance: from remote enhancers to 3D regulatory ensembles. Semin. Cell Dev. Biol. 57, 57–67 (2016).
Smith, E. & Shilatifard, A. Enhancer biology and enhanceropathies. Nat. Struct. Mol. Biol. 21, 210–219 (2014).
Bickmore, W. A. & van Steensel, B. Genome architecture: domain organization of interphase chromosomes. Cell 152, 1270–1284 (2013).
Bonev, B. & Cavalli, G. Organization and function of the 3D genome. Nat. Rev. Genet. 17, 661–678 (2016).
Rowley, M. J. & Corces, V. G. Organizational principles of 3D genome architecture. Nat. Rev. Genet. 19, 789–800 (2018).
Schmitt, A. D., Hu, M. & Ren, B. Genome-wide mapping and analysis of chromosome architecture. Nat. Rev. Mol. Cell Biol. 17, 743–755 (2016).
Creyghton, M. P. et al. Histone H3K27ac separates active from poised enhancers and predicts developmental state. Proc. Natl Acad. Sci. USA 107, 21931–21936 (2010).
Rada-Iglesias, A. et al. A unique chromatin signature uncovers early developmental enhancers in humans. Nature 470, 279–283 (2011).
Zentner, G. E., Tesar, P. J. & Scacheri, P. C. Epigenetic signatures distinguish multiple classes of enhancers with distinct cellular functions. Genome Res. 21, 1273–1283 (2011).
Carter, D. et al. Long-range chromatin regulatory interactions in vivo. Nat. Genet. 32, 623–626 (2002).
Morgan, S. L. et al. Manipulation of nuclear architecture through CRISPR-mediated chromosomal looping. Nat. Commun. 8, 15993 (2017).
Rege, M. et al. LADL: light-activated dynamic looping for endogenous gene expression control. Preprint at bioRxiv https://www.biorxiv.org/content/10.1101/349340v2 (2018).
Chubb, J. R., Trcek, T., Shenoy, S. M. & Singer, R. H. Transcriptional pulsing of a developmental gene. Curr. Biol. 16, 1018–1025 (2006).
Raj, A., Peskin, C. S., Tranchina, D., Vargas, D. Y. & Tyagi, S. Stochastic mRNA synthesis in mammalian cells. PLOS Biol. 4, e309 (2006).
Lee, K., Hsiung, C. C., Huang, P., Raj, A. & Blobel, G. A. Dynamic enhancer-gene body contacts during transcription elongation. Genes Dev. 29, 1992–1997 (2015).
Mitchell, J. A. & Fraser, P. Transcription factories are nuclear subcompartments that remain in the absence of transcription. Genes Dev. 22, 20–25 (2008).
Papantonis, A. et al. Active RNA polymerases: mobile or immobile molecular machines? PLOS Biol. 8, e1000419 (2010).
Benabdallah, N. S. et al. PARP mediated chromatin unfolding is coupled to long-range enhancer activation. Preprint at bioRxiv https://www.biorxiv.org/content/10.1101/155325v1 (2017).
Gupta, R. M. et al. A genetic variant associated with five vascular diseases is a distal regulator of endothelin-1 gene expression. Cell 170, 522–533 (2017).
Bonev, B. et al. Multiscale 3D genome rewiring during mouse neural development. Cell 171, 557–572 (2017).
Chen, H. et al. Dynamic interplay between enhancer-promoter topology and gene activity. Nat. Genet. 50, 1296–1303 (2018). In this study, simultaneous live imaging of long-range enhancer–promoter contacts and transcriptional output demonstrate that sustained physical proximity between the enhancer and its target promoter is required for gene activation.
Jeong, Y., El-Jaick, K., Roessler, E., Muenke, M. & Epstein, D. J. A functional screen for sonic hedgehog regulatory elements across a 1 Mb interval identifies long-range ventral forebrain enhancers. Development 133, 761–772 (2006).
Sagai, T. et al. A cluster of three long-range enhancers directs regional Shh expression in the epithelial linings. Development 136, 1665–1674 (2009).
Dixon, J. R. et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485, 376–380 (2012).
Jin, F. et al. A high-resolution map of the three-dimensional chromatin interactome in human cells. Nature 503, 290–294 (2013).
Williamson, I., Lettice, L. A., Hill, R. E. & Bickmore, W. A. Shh and ZRS enhancer colocalisation is specific to the zone of polarising activity. Development 143, 2994–3001 (2016).
Sagai, T., Hosoya, M., Mizushina, Y., Tamura, M. & Shiroishi, T. Elimination of a long-range cis-regulatory module causes complete loss of limb-specific Shh expression and truncation of the mouse limb. Development 132, 797–803 (2005).
Kvon, E. Z. et al. Progressive loss of function in a limb enhancer during snake evolution. Cell 167, 633–642 (2016).
Lettice, L. A., Hill, A. E., Devenney, P. S. & Hill, R. E. Point mutations in a distant sonic hedgehog cis-regulator generate a variable regulatory output responsible for preaxial polydactyly. Hum. Mol. Genet. 17, 978–985 (2008).
Lieberman-Aiden, E. et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009). This work established Hi-C as a method to capture chromosomal interactions genome-wide.
Nora, E. P. et al. Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 485, 381–385 (2012).
Sexton, T. et al. Three-dimensional folding and functional organization principles of the Drosophila genome. Cell 148, 458–472 (2012).
Dowen, J. M. et al. Control of cell identity genes occurs in insulated neighborhoods in mammalian chromosomes. Cell 159, 374–387 (2014).
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).
Tang, Z. et al. CTCF-mediated human 3D genome architecture reveals chromatin topology for transcription. Cell 163, 1611–1627 (2015).
Hnisz, D., Day, D. S. & Young, R. A. Insulated neighborhoods: structural and functional units of mammalian gene control. Cell 167, 1188–1200 (2016).
Dixon, J. R. et al. Chromatin architecture reorganization during stem cell differentiation. Nature 518, 331–336 (2015).
Symmons, O. et al. Functional and topological characteristics of mammalian regulatory domains. Genome Res. 24, 390–400 (2014).
Symmons, O. et al. The Shh topological domain facilitates the action of remote enhancers by reducing the effects of genomic distances. Dev. Cell 39, 529–543 (2016). This paper shows that engineered chromosomal rearrangements support a role for the Shh TADs as a structural facilitator of regulatory long-range interactions.
Schoenfelder, S. et al. Polycomb repressive complex PRC1 spatially constrains the mouse embryonic stem cell genome. Nat. Genet. 47, 1179–1186 (2015).
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).
Hnisz, D. et al. Activation of proto-oncogenes by disruption of chromosome neighborhoods. Science 351, 1454–1458 (2016).
Ji, X. et al. 3D chromosome regulatory landscape of human pluripotent cells. Cell Stem Cell 18, 262–275 (2016).
Liu, X. S. et al. Editing DNA methylation in the mammalian genome. Cell 167, 233–247 (2016).
Le Dily, F. et al. Distinct structural transitions of chromatin topological domains correlate with coordinated hormone-induced gene regulation. Genes Dev. 28, 2151–2162 (2014).
Bintu, B. et al. Super-resolution chromatin tracing reveals domains and cooperative interactions in single cells. Science 362, eaau1783 (2018). In this study, super-resolution microscopy of the chromatin structure uncovers TAD-like domains in single cells that persist in the absence of cohesin.
Phillips-Cremins, J. E. et al. Architectural protein subclasses shape 3D organization of genomes during lineage commitment. Cell 153, 1281–1295 (2013).
Caputo, L. et al. The Isl1/Ldb1 complex orchestrates genome-wide chromatin organization to instruct differentiation of multipotent cardiac progenitors. Cell Stem Cell 17, 287–299 (2015).
Krivega, I. & Dean, A. LDB1-mediated enhancer looping can be established independent of mediator and cohesin. Nucleic Acids Res. 45, 8255–8268 (2017).
Nasmyth, K. Disseminating the genome: joining, resolving, and separating sister chromatids during mitosis and meiosis. Annu. Rev. Genet. 35, 673–745 (2001).
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).
Parelho, V. et al. Cohesins functionally associate with CTCF on mammalian chromosome arms. Cell 132, 422–433 (2008).
Wendt, K. S. et al. Cohesin mediates transcriptional insulation by CCCTC-binding factor. Nature 451, 796–801 (2008).
Murrell, A., Heeson, S. & Reik, W. Interaction between differentially methylated regions partitions the imprinted genes Igf2 and H19 into parent-specific chromatin loops. Nat. Genet. 36, 889–893 (2004).
Kurukuti, S. et al. CTCF binding at the H19 imprinting control region mediates maternally inherited higher-order chromatin conformation to restrict enhancer access to Igf2. Proc. Natl Acad. Sci. USA 103, 10684–10689 (2006).
Narendra, V. et al. CTCF establishes discrete functional chromatin domains at the Hox clusters during differentiation. Science 347, 1017–1021 (2015).
Guo, Y. et al. CRISPR inversion of CTCF sites alters genome topology and enhancer/promoter function. Cell 162, 900–910 (2015).
de Wit, E. et al. CTCF binding polarity determines chromatin looping. Mol. Cell 60, 676–684 (2015).
Hanssen, L. L. P. et al. Tissue-specific CTCF-cohesin-mediated chromatin architecture delimits enhancer interactions and function in vivo. Nat. Cell Biol. 19, 952–961 (2017).
Cuartero, S. et al. Control of inducible gene expression links cohesin to hematopoietic progenitor self-renewal and differentiation. Nat. Immunol. 19, 932–941 (2018).
Schmidt, D. et al. A CTCF-independent role for cohesin in tissue-specific transcription. Genome Res. 20, 578–588 (2010).
Kagey, M. H. et al. Mediator and cohesin connect gene expression and chromatin architecture. Nature 467, 430–435 (2010).
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).
Bracken, A. P. & Helin, K. Polycomb group proteins: navigators of lineage pathways led astray in cancer. Nat. Rev. Cancer 9, 773–784 (2009).
Denholtz, M. et al. Long-range chromatin contacts in embryonic stem cells reveal a role for pluripotency factors and Polycomb proteins in genome organization. Cell Stem Cell 13, 602–616 (2013).
Isoda, T. et al. Non-coding transcription instructs chromatin folding and compartmentalization to dictate enhancer-promoter communication and T cell fate. Cell 171, 103–119 (2017).
Tsai, P. F. et al. A muscle-specific enhancer RNA mediates cohesin recruitment and regulates transcription in trans. Mol. Cell 71, 129–141 (2018).
Engreitz, J. M. et al. The Xist lncRNA exploits three-dimensional genome architecture to spread across the X chromosome. Science 341, 1237973 (2013).
Cajigas, I. et al. The Evf2 ultraconserved enhancer lncRNA functionally and spatially organizes megabase distant genes in the developing forebrain. Mol. Cell 71, 956–972 (2018).
Perry, R. B., Hezroni, H., Goldrich, M. J. & Ulitsky, I. Regulation of neurodegeneration by long noncoding RNAs. Mol. Cell 72, 553–567 (2018).
Anderson, K. M. et al. Transcription of the non-coding RNA upperhand controls Hand2 expression and heart development. Nature 539, 433–436 (2016).
Hacisuleyman, E. et al. Topological organization of multichromosomal regions by the long intergenic noncoding RNA Firre. Nat. Struct. Mol. Biol. 21, 198–206 (2014).
Lai, F. et al. Activating RNAs associate with Mediator to enhance chromatin architecture and transcription. Nature 494, 497–501 (2013).
Kung, J. T. et al. Locus-specific targeting to the X chromosome revealed by the RNA interactome of CTCF. Mol. Cell 57, 361–375 (2015).
Sigova, A. A. et al. Transcription factor trapping by RNA in gene regulatory elements. Science 350, 978–981 (2015).
Busslinger, G. A. et al. Cohesin is positioned in mammalian genomes by transcription, CTCF and Wapl. Nature 544, 503–507 (2017).
Heinz, S. et al. Transcription elongation can affect genome 3D structure. Cell 174, 1522–1536 (2018).
Gu, B. et al. Transcription-coupled changes in nuclear mobility of mammalian cis-regulatory elements. Science 359, 1050–1055 (2018). This study uses live imaging of regulatory elements to reveal a transcription-dependent increased mobility of both enhancers and promoters upon activation.
Osborne, C. S. et al. Active genes dynamically colocalize to shared sites of ongoing transcription. Nat. Genet. 36, 1065–1071 (2004).
de Laat, W. & Duboule, D. Topology of mammalian developmental enhancers and their regulatory landscapes. Nature 502, 499–506 (2013).
Drissen, R. et al. The active spatial organization of the β-globin locus requires the transcription factor EKLF. Genes Dev. 18, 2485–2490 (2004).
Vernimmen, D., De Gobbi, M., Sloane-Stanley, J. A., Wood, W. G. & Higgs, D. R. Long-range chromosomal interactions regulate the timing of the transition between poised and active gene expression. EMBO J. 26, 2041–2051 (2007).
Montavon, T. et al. A regulatory archipelago controls Hox genes transcription in digits. Cell 147, 1132–1145 (2011).
van de Werken, H. J. et al. Robust 4C-seq data analysis to screen for regulatory DNA interactions. Nat. Methods 9, 969–972 (2012).
Kim, Y. H. et al. Rev-erbα dynamically modulates chromatin looping to control circadian gene transcription. Science 359, 1274–1277 (2018).
Mermet, J. et al. Clock-dependent chromatin topology modulates circadian transcription and behavior. Genes Dev. 32, 347–358 (2018).
Comoglio, F. et al. Thrombopoietin signaling to chromatin elicits rapid and pervasive epigenome remodeling within poised chromatin architectures. Genome Res. 28, 295–309 (2018).
Melo, C. A. et al. eRNAs are required for p53-dependent enhancer activity and gene transcription. Mol. Cell 49, 524–535 (2013).
Cruz-Molina, S. et al. PRC2 facilitates the regulatory topology required for poised enhancer function during pluripotent stem cell differentiation. Cell Stem Cell 20, 689–705 (2017).
Ghavi-Helm, Y. et al. Enhancer loops appear stable during development and are associated with paused polymerase. Nature 512, 96–100 (2014).
Herranz, D. et al. A NOTCH1-driven MYC enhancer promotes T cell development, transformation and acute lymphoblastic leukemia. Nat. Med. 20, 1130–1137 (2014).
Bauer, D. E. et al. An erythroid enhancer of BCL11A subject to genetic variation determines fetal hemoglobin level. Science 342, 253–257 (2013).
Smemo, S. et al. Obesity-associated variants within FTO form long-range functional connections with IRX3. Nature 507, 371–375 (2014).
Soldner, F. et al. Parkinson-associated risk variant in distal enhancer of α-synuclein modulates target gene expression. Nature 533, 95–99 (2016).
Maurano, M. T. et al. Systematic localization of common disease-associated variation in regulatory DNA. Science 337, 1190–1195 (2012).
Davison, L. J. et al. Long-range DNA looping and gene expression analyses identify DEXI as an autoimmune disease candidate gene. Hum. Mol. Genet. 21, 322–333 (2012).
Dryden, N. H. et al. Unbiased analysis of potential targets of breast cancer susceptibility loci by capture Hi-C. Genome Res. 24, 1854–1868 (2014).
Jager, R. et al. Capture Hi-C identifies the chromatin interactome of colorectal cancer risk loci. Nat. Commun. 6, 6178 (2015).
Martin, P. et al. Capture Hi-C reveals novel candidate genes and complex long-range interactions with related autoimmune risk loci. Nat. Commun. 6, 10069 (2015).
McGovern, A. et al. Capture Hi-C identifies a novel causal gene, IL20RA, in the pan-autoimmune genetic susceptibility region 6q23. Genome Biol. 17, 212 (2016).
Mumbach, M. R. et al. Enhancer connectome in primary human cells identifies target genes of disease-associated DNA elements. Nat. Genet. 49, 1602–1612 (2017).
Tak, Y. G. et al. Effects on the transcriptome upon deletion of a distal element cannot be predicted by the size of the H3K27Ac peak in human cells. Nucleic Acids Res. 44, 4123–4133 (2016).
Luo, Z. et al. A prostate cancer risk element functions as a repressive loop that regulates HOXA13. Cell Rep. 21, 1411–1417 (2017).
Osterwalder, M. et al. Enhancer redundancy provides phenotypic robustness in mammalian development. Nature 554, 239–243 (2018). This paper shows that multiple enhancers with similar activities act on target genes to confer robustness during mouse limb development.
Heinz, S. et al. Effect of natural genetic variation on enhancer selection and function. Nature 503, 487–492 (2013).
Dorsett, D. & Krantz, I. D. On the molecular etiology of Cornelia de Lange syndrome. Ann. NY Acad. Sci. 1151, 22–37 (2009).
Olley, G. et al. BRD4 interacts with NIPBL and BRD4 is mutated in a Cornelia de Lange-like syndrome. Nat. Genet. 50, 329–332 (2018).
Roelfsema, J. H. et al. Genetic heterogeneity in Rubinstein-Taybi syndrome: mutations in both the CBP and EP300 genes cause disease. Am. J. Hum. Genet. 76, 572–580 (2005).
Flavahan, W. A. et al. Insulator dysfunction and oncogene activation in IDH mutant gliomas. Nature 529, 110–114 (2016).
Groschel, S. et al. A single oncogenic enhancer rearrangement causes concomitant EVI1 and GATA2 deregulation in leukemia. Cell 157, 369–381 (2014).
Ing-Simmons, E. et al. Spatial enhancer clustering and regulation of enhancer-proximal genes by cohesin. Genome Res. 25, 504–513 (2015).
Beagrie, R. A. et al. Complex multi-enhancer contacts captured by genome architecture mapping. Nature 543, 519–524 (2017).
Markenscoff-Papadimitriou, E. et al. Enhancer interaction networks as a means for singular olfactory receptor expression. Cell 159, 543–557 (2014).
Schoenfelder, S. et al. Preferential associations between co-regulated genes reveal a transcriptional interactome in erythroid cells. Nat. Genet. 42, 53–61 (2010).
Fanucchi, S., Shibayama, Y., Burd, S., Weinberg, M. S. & Mhlanga, M. M. Chromosomal contact permits transcription between coregulated genes. Cell 155, 606–620 (2013).
Zhao, Z. et al. Circular chromosome conformation capture (4C) uncovers extensive networks of epigenetically regulated intra- and interchromosomal interactions. Nat. Genet. 38, 1341–1347 (2006).
Apostolou, E. & Thanos, D. Virus infection induces NF-κB-dependent interchromosomal associations mediating monoallelic IFN-β gene expression. Cell 134, 85–96 (2008).
Monahan, K., Horta, A. & Lomvardas, S. LHX2- and LDB1-mediated trans interactions regulate olfactory receptor choice. Nature 565, 448–453 (2019). This paper uncovers the enhancer-driven assembly of a trans -chromosomal interaction hub involving olfactory receptor genes and their regulatory elements, which mediates monoallelic olfactory receptor gene expression in mouse sensory neurons.
Noordermeer, D. et al. Variegated gene expression caused by cell-specific long-range DNA interactions. Nat. Cell Biol. 13, 944–951 (2011).
Johnston, R. J. & Desplan, C. Interchromosomal communication coordinates intrinsically stochastic expression between alleles. Science 343, 661–665 (2014).
Grubert, F. et al. Genetic control of chromatin states in humans involves local and distal chromosomal interactions. Cell 162, 1051–1065 (2015).
Waszak, S. M. et al. Population variation and genetic control of modular chromatin architecture in humans. Cell 162, 1039–1050 (2015).
Visel, A. et al. ChIP-seq accurately predicts tissue-specific activity of enhancers. Nature 457, 854–858 (2009).
Visel, A., Minovitsky, S., Dubchak, I. & Pennacchio, L. A. VISTA Enhancer Browser — a database of tissue-specific human enhancers. Nucleic Acids Res. 35, D88–D92 (2007).
Barakat, T. S. et al. Functional dissection of the enhancer repertoire in human embryonic stem cells. Cell Stem Cell 23, 276–288 (2018).
Pradeepa, M. M. et al. Histone H3 globular domain acetylation identifies a new class of enhancers. Nat. Genet. 48, 681–686 (2016).
Andersson, R., Sandelin, A. & Danko, C. G. A unified architecture of transcriptional regulatory elements. Trends Genet. 31, 426–433 (2015).
Xu, Z., Wei, G., Chepelev, I., Zhao, K. & Felsenfeld, G. Mapping of INS promoter interactions reveals its role in long-range regulation of SYT8 transcription. Nat. Struct. Mol. Biol. 18, 372–378 (2011).
Dao, L. T. M. et al. Genome-wide characterization of mammalian promoters with distal enhancer functions. Nat. Genet. 49, 1073–1081 (2017).
Akhtar, W. et al. Chromatin position effects assayed by thousands of reporters integrated in parallel. Cell 154, 914–927 (2013).
Cairns, J. et al. CHiCAGO: robust detection of DNA looping interactions in capture Hi-C data. Genome Biol. 17, 127 (2016).
Allahyar, A. et al. Enhancer hubs and loop collisions identified from single-allele topologies. Nat. Genet. 50, 1151–1160 (2018). This study uses long-read sequencing to reveal contacts involving multiple regulatory elements at the level of individual alleles.
Fukaya, T., Lim, B. & Levine, M. Enhancer control of transcriptional bursting. Cell 166, 358–368 (2016).
Hong, J. W., Hendrix, D. A. & Levine, M. S. Shadow enhancers as a source of evolutionary novelty. Science 321, 1314 (2008).
Frankel, N. et al. Phenotypic robustness conferred by apparently redundant transcriptional enhancers. Nature 466, 490–493 (2010).
Zhou, H. Y. et al. A Sox2 distal enhancer cluster regulates embryonic stem cell differentiation potential. Genes Dev. 28, 2699–2711 (2014).
Moorthy, S. D. et al. Enhancers and super-enhancers have an equivalent regulatory role in embryonic stem cells through regulation of single or multiple genes. Genome Res. 27, 246–258 (2017).
Bahr, C. et al. A Myc enhancer cluster regulates normal and leukaemic haematopoietic stem cell hierarchies. Nature 553, 515–520 (2018).
Gonen, N. et al. Sex reversal following deletion of a single distal enhancer of Sox9. Science 360, 1469–1473 (2018).
Gonen, N., Quinn, A., O’Neill, H. C., Koopman, P. & Lovell-Badge, R. Normal levels of Sox9 expression in the developing mouse testis depend on the TES/TESCO enhancer, but this does not act alone. PLOS Genet. 13, e1006520 (2017).
Ragoczy, T. et al. The locus control region is required for association of the murine β-globin locus with engaged transcription factories during erythroid maturation. Genes Dev. 20, 1447–1457 (2006).
Hyman, A. A. & Simons, K. Beyond oil and water — phase transitions in cells. Science 337, 1047–1049 (2012).
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).
Cho, W. K. et al. Mediator and RNA polymerase II clusters associate in transcription-dependent condensates. Science 361, 412–415 (2018).
Chong, S. et al. Imaging dynamic and selective low-complexity domain interactions that control gene transcription. Science 361, eaar2555 (2018).
Sabari, B. R. et al. Coactivator condensation at super-enhancers links phase separation and gene control. Science 361, eaar3958 (2018).
Boija, A. et al. Transcription factors activate genes through the phase-separation capacity of their activation domains. Cell 175, 1842–1855 (2018).
Saha, S. et al. Polar positioning of phase-separated liquid compartments in cells regulated by an mRNA competition mechanism. Cell 166, 1572–1584 (2016).
Zabidi, M. A. et al. Enhancer-core-promoter specificity separates developmental and housekeeping gene regulation. Nature 518, 556–559 (2015).
Nagano, T. et al. Single-cell Hi-C reveals cell-to-cell variability in chromosome structure. Nature 502, 59–64 (2013).
Nagano, T. et al. Cell-cycle dynamics of chromosomal organization at single-cell resolution. Nature 547, 61–67 (2017).
Dekker, J., Rippe, K., Dekker, M. & Kleckner, N. Capturing chromosome conformation. Science 295, 1306–1311 (2002).
Simonis, M. et al. Nuclear organization of active and inactive chromatin domains uncovered by chromosome conformation capture-on-chip (4C). Nat. Genet. 38, 1348–1354 (2006).
Dostie, J. et al. Chromosome conformation capture carbon copy (5C): a massively parallel solution for mapping interactions between genomic elements. Genome Res. 16, 1299–1309 (2006).
Belton, J. M. et al. Hi-C: a comprehensive technique to capture the conformation of genomes. Methods 58, 268–276 (2012).
Hughes, J. R. et al. Analysis of hundreds of cis-regulatory landscapes at high resolution in a single, high-throughput experiment. Nat. Genet. 46, 205–212 (2014).
Sahlen, P. et al. Genome-wide mapping of promoter-anchored interactions with close to single-enhancer resolution. Genome Biol. 16, 156 (2015).
Fullwood, M. J. et al. An oestrogen-receptor-α-bound human chromatin interactome. Nature 462, 58–64 (2009).
Mumbach, M. R. et al. HiChIP: efficient and sensitive analysis of protein-directed genome architecture. Nat. Methods 13, 919–922 (2016).
Brant, L. et al. Exploiting native forces to capture chromosome conformation in mammalian cell nuclei. Mol. Syst. Biol. 12, 891 (2016).
Williamson, I. et al. Spatial genome organization: contrasting views from chromosome conformation capture and fluorescence in situ hybridization. Genes Dev. 28, 2778–2791 (2014).
Boettiger, A. N. et al. Super-resolution imaging reveals distinct chromatin folding for different epigenetic states. Nature 529, 418–422 (2016).
Arnold, C. D. et al. Genome-wide quantitative enhancer activity maps identified by STARR-seq. Science 339, 1074–1077 (2013).
Dickel, D. E. et al. Function-based identification of mammalian enhancers using site-specific integration. Nat. Methods 11, 566–571 (2014).
Canver, M. C. et al. BCL11A enhancer dissection by Cas9-mediated in situ saturating mutagenesis. Nature 527, 192–197 (2015).
Korkmaz, G. et al. Functional genetic screens for enhancer elements in the human genome using CRISPR-Cas9. Nat. Biotechnol. 34, 192–198 (2016).
Thakore, P. I. et al. Highly specific epigenome editing by CRISPR-Cas9 repressors for silencing of distal regulatory elements. Nat. Methods 12, 1143–1149 (2015).
Kearns, N. A. et al. Functional annotation of native enhancers with a Cas9-histone demethylase fusion. Nat. Methods 12, 401–403 (2015).
Fulco, C. P. et al. Activity-by-contact model of enhancer specificity from thousands of CRISPR perturbations. Preprint at bioRxiv https://www.biorxiv.org/content/10.1101/529990v1 (2019).
Gasperini, M. et al. A genome-wide framework for mapping gene regulation via cellular genetic screens. Cell 176, 377–390 (2019).
Hilton, I. B. et al. Epigenome editing by a CRISPR-Cas9-based acetyltransferase activates genes from promoters and enhancers. Nat. Biotechnol. 33, 510–517 (2015).
Acknowledgements
The authors thank L. Chakalova, N. Gonen and M. Osterwalder for insightful comments on the manuscript, and E. Nora for valuable discussions. They apologize to colleagues whose important primary studies they were unable to cite due to space constraints. P.F. was supported by grants from the Medical Research Council UK (MR/L007150/1), the US National Institutes of Health (4D Nucleome 1U01HL129971-01), the European Research Council (340152) and the Biotechnology and Biological Science Research Council UK (BB/J004480/1). S.S. was supported by the Biotechnology and Biological Science Research Council UK (BB/J004480/1) and a Career Progression Fellowship from the Babraham Institute.
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Glossary
- dCas9
-
A nuclease-deficient variant of the Cas9 protein. As for Cas9, it can be targeted to defined genomic loci by a guide RNA. dCas9 is commonly used as a fusion protein, thus allowing locus-specific recruitment of domains conferring activities such as transcriptional modulation, chromatin modification or intermolecular dimerization.
- Transcription factories
-
Discrete nuclear foci of concentrated, hyperphosphorylated RNA polymerase II at which nascent transcription occurs. Active genes and their regulatory elements have been shown to coalesce at transcription factories, and the dynamic relocation of genes towards and away from transcription factories correlates with their transcriptional ‘on’ and ‘off’ cycles.
- Super-enhancer
-
A class of regulatory regions in mammalian genomes with an unusually high degree of enrichment for the binding of the transcriptional co-activator mediator complex subunit 1 (MED1), the histone mark H3K27ac or cell-type-specific transcription factors. As they often result from concatenating individual enhancers, super-enhancers on average span considerably larger genomic regions than ‘normal’ enhancers. Super-enhancers are often located in the proximity of lineage-specifying genes, which they have been proposed to control. The extent to which super-enhancers represent a class of regulatory elements that is functionally distinct from ‘normal’ enhancers is under debate.
- Expression quantitative trait loci
-
(eQTLs). Genomic loci in which genetic variants can be causally linked to variation of gene expression levels, either in cis or, more rarely, in trans. eQTLs are identified by testing the linkage between variation in expression and genetic polymorphisms. eQTL mapping has become a widely used tool to identify genetic sequence variants that affect gene expression control.
- TRIP
-
Thousands of reporters integrated in parallel (TRIP) is a high-throughput assay based on the random integration of barcoded GFP reporter genes (under the control of a minimal promoter) into the genome, to reveal the effects of cis-regulatory elements and the local chromatin environment on gene expression.
- Shadow enhancers
-
Discovered in Drosophila melanogaster, these ‘secondary’ enhancers appear to act in a largely redundant manner to ‘primary’ enhancers; that is, both enhancers drive similar patterns of expression, and deletion of either enhancer does not result in aberrant phenotypes. The largely overlapping functions of primary and secondary enhancers serve to buffer gene expression patterns against genetic or environmental perturbations, thereby conferring robustness during development.
- Pioneer transcription factors
-
A class of transcription factors that can bind to target sites in compacted (‘closed’) chromatin, which is covered by nucleosomes and is not DNase I hypersensitive. Pioneer transcription factors are thought to remodel the chromatin landscape during early steps of cell fate decisions to facilitate the subsequent recruitment of other (non-pioneer) transcription factors and DNA-modifying and chromatin-modifying enzymes to their target sites.
- Phase-separated domains
-
Distinct subcellular non-membrane-bound domains, in the nucleus or in the cytoplasm. They are dynamic compartments in which the components are concentrated relative to the surroundings. Phase-separated condensates in the nucleus have been implicated in various aspects of genome control, including gene expression and heterochromatin formation.
- Low-complexity domains
-
Protein domains (also known as intrinsically disordered regions) that are often found in the activation domains of transcription factors. They contain repeats of single amino acids or short amino acid motifs that are usually not amenable to crystallography. Interactions between low-complexity domains promote the aggregation of transcription factors and cofactors in droplets in vitro and are required for the formation of phase-separated condensates in the nucleus, which have been proposed to drive transcriptional control.
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Schoenfelder, S., Fraser, P. Long-range enhancer–promoter contacts in gene expression control. Nat Rev Genet 20, 437–455 (2019). https://doi.org/10.1038/s41576-019-0128-0
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DOI: https://doi.org/10.1038/s41576-019-0128-0
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