Imaging studies, high-resolution chromatin conformation maps, and genome-wide occupancy data of architectural proteins have revealed that genome topology is tightly intertwined with gene expression. Cross-talk between gene-regulatory elements is often organized within insulated neighborhoods, and regulatory cues that induce transcriptional changes can reshape chromatin folding patterns and gene positioning within the nucleus. The cause–consequence relationship of genome architecture and gene expression is intricate, and its molecular mechanisms are under intense investigation. Here, we review the interdependency of transcription and genome organization with emphasis on enhancer–promoter contacts in gene regulation.

Overview

The genetic material of eukaryotic cells is encased by the nuclear envelope. The inner of the two nuclear membranes is attached to a fibrillar network of proteins that constitutes the nuclear lamina, which has a role in genome organization (Akhtar and Gasser, 2007; van Steensel and Belmont, 2017). Chromosomes occupy distinct territories within the nucleus, and their radial positioning correlates with gene content and activity (Cremer et al., 2006). Gene poor regions and repressed genes are closer to the nuclear periphery and are often found in close contact with the lamina as part of lamina-associated domains (van Steensel and Belmont, 2017). Gene-rich, active regions tend to localize centrally (Bickmore and van Steensel, 2013) but are also found at nuclear pore complexes that perforate the nuclear envelope. In addition, the nucleoplasm contains numerous presumably self-organizing structures with roles in gene expression, such as the nucleolus, Cajal bodies, splicing bodies, and promyelocytic leukemia bodies, that are not surrounded by membranes (Mao et al., 2011). Transcriptionally inactive chromatin can aggregate near nuclear chromocenters (pericentromere-associated domains; Wijchers et al., 2015) and is found near the nucleolus in so-called nucleolus associating domains (Németh et al., 2010; van Koningsbruggen et al., 2010). Repositioning of loci from the nucleolus to the lamina after cell division suggests mobility of transcriptionally repressed DNA between different heterochromatic sites (Kind et al., 2013). Differentiation-induced activation can move genes away from the lamina (Peric-Hupkes et al., 2010), and as an additional example of gene mobility upon activation, loci can be extruded from chromosome territories (CTs; Ragoczy et al., 2003; Chambeyron and Bickmore, 2004). This indicates that gene positioning within different nuclear environments is closely linked to gene activity (Fig. 1).

Imaging studies have provided numerous fundamental insights into global nuclear organization, but techniques based on chromosome conformation capture (3C) enabled unprecedented views of chromosome folding, reaching single-kilobase resolution of chromatin contacts (Cullen et al., 1993; Dekker et al., 2002). 3C and its high-throughput derivatives (Denker and de Laat, 2016) are based on proximity ligation of DNA fragments and have detected numerous layers of chromosome organization. At the lowest scale, transcriptional regulation was found to involve long-range contacts between regulatory elements, such as enhancers, and their target genes. Enhancers serve as a binding platform for transcription factors that can boost gene transcription, and a characteristic chromatin signature has allowed for their annotation in many cell types (Calo and Wysocka, 2013). Enhancers mostly reside in the noncoding part of the genome and they often act in a cell type–specific manner, thereby establishing specialized gene expression programs characteristic of specific cell types. Genes can be influenced by multiple enhancers that can be located up to over a megabase away from their target promoter, but in many cases, chromatin looping was found to allow for communication between elements regardless of distance. However, the regulatory space of enhancers is confined by megabase scale topologically associating domains (TADs) that are flanked by boundary elements across which the probability of chromatin interactions is reduced (Dixon et al., 2012; Nora et al., 2012; Sexton et al., 2012). The tendency of active and repressive chromatin to segregate and occupy distinct regions in the nucleus is referred to as A (transcriptionally active) and B (inactive) compartments (Lieberman-Aiden et al., 2009). In addition, intra-chromosomal contacts are more frequent than inter-chromosomal contacts, which is compatible with the concept of CTs that had been described in imaging studies (Cremer and Cremer, 2001).

In this review we explore in greater depth the mutual relationship between enhancer function and architectural features of chromatin, including how certain interactions are favored and stabilized while others are disfavored, and how enhancer activity is regulated by and impacts upon a complex, multi-layered nuclear architectural framework.

Physical proximity can mediate communication among regulatory elements

Transcriptional enhancers are DNA sequences that augment gene transcription (Banerji et al., 1981; Moreau et al., 1981). Enhancers function via a plethora of mechanisms that are initiated by sequence-specific DNA binding proteins and their coregulators (Bulger and Groudine, 2011; Buecker and Wysocka, 2012; de Laat and Duboule, 2013). Enhancer-bound transcription factors can impact gene activity by initiating the opening of chromatin via recruitment of chromatin remodeling complexes to generate nucleosome free regions. They can also recruit histone modifying complexes, basal transcription factors including RNA polymerase 2 (pol2), and/or proteins involved in transcriptional pause-release. Many of these features along with sequence conservation have been used to annotate enhancers genome-wide (Calo and Wysocka, 2013).

Enhancers typically contain clusters of transcription factor binding sites (Fig. 1). Transcription factors and their cofactors can form assemblies (termed enhanceosomes) via cooperative chromatin binding, which can result in synergistic transcriptional outcomes (Merika and Thanos, 2001). In addition, genes can be controlled by multiple enhancers that in some cases are close to each other along the linear chromosome (Fig. 1). Clusters of enhancer elements have been varyingly named locus control region (LCR) as well as stretch-, spread-, or super-enhancers based on size, histone modifications, and level of occupancy by nuclear factors (Heinz et al., 2015; Pott and Lieb, 2015). Constituent enhancer elements can also be scattered across a locus, residing upstream, within, or downstream of the genes they control, and can cluster in 3D to form “hubs” (Patrinos et al., 2004). Yet neither the size of a composite enhancer nor its positioning allows for reliable predictions of its mechanism of action.

Vertebrate enhancers can be located up to over a megabase away from the promoters they act on (Lettice et al., 2003; Sagai et al., 2004). An answer to how these elements can communicate over large distances came with the development of 3C (Cullen et al., 1993; Dekker et al., 2002). 3C and its high-throughput derivatives (4C, 5C, capture-C, and Hi-C, among others) employ stabilization of chromosomal contacts via chemical cross-linking, restriction digestion of DNA, and ligation of DNA fragments, followed by quantification of chimeric DNA fragments (Denker and de Laat, 2016). The underlying concept is that the closer genomic regions are positioned in nuclear space, the more likely they are to be cross-linked and ligated to each other. Original analyses of the β-globin locus by 3C (Tolhuis et al., 2002) or a method called RNA-TRAP (Carter et al., 2002) revealed that an enhancer, the LCR, contacts the β-globin promoter, looping out the intervening ∼50 kb of DNA. The β-globin LCR, which is used frequently here as an example, is a strong erythroid-specific enhancer required for the normal expression of all genes in the cluster. It consists of a cluster of DNase I hypersensitive sites that cooperate to ensure proper globin expression during erythroid differentiation (Bender et al., 1998, 2001; Bulger et al., 2003; Fang et al., 2005). Similar regulatory loops were found for other genes (e.g., Murrell et al., 2004a; Würtele and Chartrand, 2006; Vernimmen et al., 2007), and broad assessments of 3D contacts have revealed that looping is a widespread phenomenon (Sanyal et al., 2012; Rao et al., 2014). Systematic examination of chromatin contacts during lineage commitment and development revealed that certain contacts are stable across different developmental stages and cell types, while others, mostly associated with variable chromatin states, are dynamic (Mifsud et al., 2015; Schoenfelder et al., 2015; Javierre et al., 2016; Andrey et al., 2017; Freire-Pritchett et al., 2017). Stable loops may make genes permissive for rapid transcriptional activation upon external stimuli but may also serve as a structure within which more dynamic contacts are formed. The fact that some enhancer–promoter interactions are established de novo to initiate gene expression while other loops are preformed indicates that contacts among regulatory elements do not necessarily result in active transcription and that expression or recruitment of additional transcription factors may be required for the loop to become functional. Moreover, ongoing transcription is not necessary to maintain enhancer–promoter loops (Mitchell and Fraser, 2008; Palstra et al., 2008).

It is important to realize that enhancer–promoter looping, even though widespread, might not be a universal mechanism of distal enhancer function. For example, a neuronal enhancer at the sonic hedgehog (Shh) locus appears to decompact the locus upon activation, thereby increasing its distance from the Shh promoter (Benabdallah et al., 2017 ,Preprint). This particular scenario seems irreconcilable with an enhancer–promoter looping mechanism. Additional experiments in this study support a mechanism by which enhancer activity spreads along the chromosome to activate the promoter. Nonetheless, distinct Shh enhancers have been shown to engage in looped contacts with the Shh gene promoter (Amano et al., 2009; Williamson et al., 2016). It is intriguing that seemingly different mechanisms evolved to control the Shh promoter over large distances in distinct cell types. An additional consideration is that proximity among regulatory elements might not necessarily be a reflection of simple loops but of more complex chromatin folding patterns in which active loci can be condensed (Williamson et al., 2016). Hence, both de-compaction and compaction (perhaps as a result of complex looping structures) can occur upon enhancer activation. Indeed, architectural studies at the Igh locus suggest that spatial confinement may help in forging enhancer–promoter interactions (Lucas et al., 2014; as will be discussed). In conclusion, transcriptional output is dependent on distal enhancer–promoter communication, which can be enabled by chromatin looping. As multiple elements or forces may be involved, high-resolution analysis (using multiple techniques/viewpoints/anchors/probes) may be necessary to interpret the contact complexity of specific loci.

Functional assessment of chromatin loops as a driving force of transcription

Evidence that chromatin loops can be instructive to activate transcription came from tethering experiments in which proximity between regulatory elements was forced in living cells (Nolis et al., 2009; Deng et al., 2012). Zinc-finger–mediated tethering of the candidate looping factor Ldb1 to the β-globin promoter led to recruitment of the LCR and transcription activation in immature erythroid precursors (Deng et al., 2012). This approach was also used to rewire the LCR with a different gene to activate its expression while reducing transcription of the enhancer-deprived gene (Deng et al., 2014). dCas9-mediated tethering of YY1 to the Etv4 promoter in mouse embryonic stem cells (mESCs) increased interaction frequency between the promoter and its enhancer and resulted in increased transcription (Weintraub et al., 2017). These studies indicate that inducing proximity between regulatory elements and promoters can causally underlie transcription.

What is the temporal relationship between transcription and architectural chromatin features? Transcription is a discontinuous process with periods of intense mRNA production (bursts) followed by longer periods of transcriptional silence (Raj and van Oudenaarden, 2008). Output can be modulated by altering burst size, reflective of the number of RNA molecules produced, and/or burst fraction, reflective of frequency and duration of the bursts. Forced juxtaposition between β-globin and the LCR leads to an increase in burst frequency but not burst size (Bartman et al., 2016). Early pioneering RNA FISH studies at this locus (Wijgerde et al., 1995), later revisited using single-molecule RNA FISH (Bartman et al., 2016), provided evidence in favor of a model in which two genes that are under the control of the same enhancer display alternating and thus mutually exclusive contacts with the enhancer. Hence, alternating bursting of these genes might be a reflection of competing enhancer–promoter loops (Fig. 2 A). These findings contrast with a study using an engineered locus in Drosophila melanogaster in which a single enhancer was capable of regulating two flanking genes in a manner such that both genes exhibited synchronous bursting behavior (Fukaya et al., 2016). The latter result suggests that looped contacts are not always mutually exclusive (Fig. 2 A), and that competition for them is not a universal mechanism for gene control at multi-gene loci. Recent live measurements of the proximity between the endogenous even-skipped (eve) enhancers and an integrated eve promoter-driven LacZ reporter gene in Drosophila embryos demonstrated a close correlation between transcriptional bursting and enhancer–promoter juxtaposition (Chen et al., 2018). Together with the aforementioned studies, this suggests that dynamic chromatin contacts can underlie bursty behavior of gene transcription. Moreover, in Chen et al. (2018), the presence of an ectopic eve promoter in the reporter construct diminished expression of the endogenous gene, presumably as a result of promoter competition.

Alternating chromatin loops have also been invoked to explain changes in gene expression over much longer time scales such as during ontogeny. For example, the developmental switch in the expression of β-type and α-type globin genes during development is thought to involve mutually exclusive interactions with the enhancers (Foley and Engel, 1992; Palstra et al., 2003; Vernimmen et al., 2007), likely controlled by developmentally regulated nuclear factors that foster specific enhancer–promoter contacts, and by developmentally dynamic architectural constraints, as will be discussed below. Competition among poised or active genes for shared enhancers can modulate gene expression and even underlie aberrant gene expression in disease. De novo formation of a transcriptional start site by a gain-of-function regulatory single-nucleotide polymorphism upstream of the human α-globin genes sequestered the distal enhancer away from the α-globin promoters (Fig. 2 B), reducing their expression and resulting in α-thalassemia (De Gobbi et al., 2006). Conversely, increased expression of the MYC proto-oncogene in cancer can result from mutations in the PVT1 promoter (Fig. 2 C), a gene in close proximity of MYC, which normally competes for the same enhancer (Cho et al., 2018). CRISPR interference based mapping of functional regulatory sequences revealed similar competitive relationships between promoters and enhancers at the MYC and GATA1 loci (Fulco et al., 2016).

Genes can be activated simply by residing within the regulatory range of an enhancer even if their expression is ostensibly irrelevant to the function of the cell in which the enhancer is active, a phenomenon called the bystander effect (Fig. 1). For example, the gene CD79b, which is transcribed and functional in B-lymphocytes, sits between the human growth hormone gene cluster and a distal LCR that drives gene expression in the pituitary gland. Transcription of CD79b is activated in the pituitary gland as a result of being close to the enhancer even though it plays no discernible role in pituitary function (Cajiao et al., 2004). As another example, the NME4 gene that resides 300 kb away from the human α-globin genes is in physical contact and under the regulatory influence of the α-globin enhancer. NME4 competes for the activity of the α-globin enhancer even though it is dispensable for erythroid cell function (Lower et al., 2009). Finally, in the forced chromatin looping experiments described above, tethering of the adult type β-globin gene to the LCR additionally activated the interspersed embryonic bH1-globin gene, presumably as a result of bringing it closer to the LCR (Deng et al., 2012), which corroborates that proximity can favor regulatory influence.

Transcriptional activity is linked to nuclear positioning

Transcriptional activation (by enhancers) often results in relocalization of genes toward the nuclear interior. Enhancers such as the β-globin LCR can drive positioning of the β-globin locus away from heterochromatin (Francastel et al., 1999) and promote movement of the locus toward the nuclear interior during erythroid cell maturation, which is accompanied by increased gene activity (Ragoczy et al., 2006). The β-globin LCR directs the locus toward foci of engaged (Ser5 phosphorylated) pol2, also referred to as transcription factories (Ragoczy et al., 2006). The function, if any, of gene movement to the nuclear center, although common and generally observed across species, is unclear. As a case in point, the CFTR gene and its neighbors migrate to the nuclear center upon activation in human but not murine cells, in which they are constitutively positioned centrally (Sadoni et al., 2008), suggesting that central nuclear localization might be permissive but not instructive for gene expression. Perhaps similarly, relocalization of β-globin away from its CT does not necessarily result in transcription but primes the region for activation (Ragoczy et al., 2003). Ectopic integration of the LCR in a different chromosome moved the integration site away from the CT and resulted in activation of some, but not all, surrounding genes (Noordermeer et al., 2008). This indicates that regulatory elements can influence nuclear localization, but does not settle the question whether it is the LCR per se, chromatin opening, or active transcription that causes positioning outside the CT. Experiments in embryonic stem cells showed that targeted chromatin decondensation at a chosen gene in the absence of transcription activation is sufficient to mobilize the locus toward the nuclear interior (Therizols et al., 2014). This suggests that chromatin state but not transcription can be a driving force in nuclear repositioning.

While many observations regarding enhancer function and nuclear architecture have been made at the β-globin locus, most concepts gleaned from this locus are likely to apply to other genes as well. However, it is important to bear in mind that considerations of enhancer function in the context of genome topology are fraught with the limitation that correlation of a particular feature and gene activity does not imply causation.

Gene silencing is an important component of transcriptional programs, and maintenance of this state is indispensable for cell fate specification. The consequences of impaired gene silencing were first observed in Drosophila in the form of severe phenotypes in mutants with derepression of developmental patterning genes such as hox genes (Lewis, 1978; Struhl, 1981; Duncan, 1982; Ingham, 1985). Polycomb group (PcG) protein complexes are now known to recognize silent genes and to maintain their inactive status through repressive epigenomic modifications such as histone 3 lysine 27 trimethylation (H3K27me3; Schwartz and Pirrotta, 2007). Interchromosomal as well as intrachromosomal interactions between PcG-silenced genes were found to underlie the aggregation of silenced chromatin into PcG bodies (Pirrotta and Li, 2012), and recent super-resolution imaging in Drosophila revealed very dense packaging of PcG chromatin (Boettiger et al., 2016). There is considerable evidence that H3K27me3 and H3K9me2/3 promote peripheral localization (Harr et al., 2015), and deposition of the former repressive mark by forced EZH2 recruitment was sufficient to induce repression and compartment (active to inactive) switching (Wijchers et al., 2016). Targeted perturbations, in which genes were tethered to specific nuclear sites, further demonstrated the direct impact of gene positioning on activity (reviewed in Deng and Blobel, 2014; Bartman and Blobel, 2015). For instance, tethering genes to the nuclear lamina resulted in repression of some but not all genes (Andrulis et al., 1998; Finlan et al., 2008; Kumaran and Spector, 2008; Reddy et al., 2008). This indicates that relocalization can drive transcriptional changes but, as discussed, regulatory elements can also drive movement of loci to compartments with a different activity status. Importantly, most of the links between subnuclear positioning and gene expression are still correlative with unexplored cause–effect relationships.

Regulatory interactions in gene repression

Heterochromatinization and loss of gene-specific looping are associated with long-term gene silencing, but architectural changes that occur at the initial stages of repression are less well understood. Especially in higher eukaryotes, studies on the link between genome organization and early gene repression are underrepresented. Polycomb response elements (PREs) in Drosophila are sufficient to recruit PcG proteins that mark regions with H3K27me3, and repress nearby genes (Simon et al., 1993; Chan et al., 1994; Wang et al., 2004). PREs, like enhancers, can be distant from their targets and are able to form long-range contacts with the genes they act on (Schwartz and Cavalli, 2017). Notably, no mammalian counterpart of PREs has been identified so far. One alternative mechanism by which polycomb can be recruited to chromatin involves noncoding RNAs that can bind to the polycomb component PRC2 (Schaaf et al., 2013; Kaneko et al., 2014; Berrozpe et al., 2017). For instance, initiation of X-chromsome inactivation by Xist results from recruiting PRC2 to Xist occupied sites. Xist spreads across the X-chromsome by scanning and occupying genomic loci that reside in spatial vicinity, which then alters chromosome structure of these sites and allows H3K27me3 to spread to new, and from there other, genomic sites (Engreitz et al., 2013).

Gene repression can be accompanied by acute loss of enhancer–promoter contacts. For example, a repressive cue leads to rapid dissocation between the enhancer and promoter at the Kit locus in maturing murine erythroblasts (Jing et al., 2008). Loss of enhancer–promoter contacts upon cell state change was observed in other cell types as well and is indispensable for shutdown of the pluripotenty program and differentiation of mESCs (Whyte et al., 2012; Respuela et al., 2016; Schnappauf et al., 2016; Bonev et al., 2017). At genes under circadian regulation, rhythmic enhancer–promoter contacts are found (Aguilar-Arnal et al., 2013; Kim et al., 2018; Mermet et al., 2018), and the transcriptional repressor Rev-erbα has been implicated in the circadian disruption of enhancer–promoter loops (Kim et al., 2018). This echoes studies in Drosophila suggesting that the transcriptional repressor Snail functions to disrupt enhancer–promoter contacts, earning its description as “anti-looping” factor (Chopra et al., 2012). However, as mentioned in other contexts above, establishing a cause–effect relationship between lost enhancer activity and the disruption of long-range contacts remains an unmet challenge.

Novel chromatin loops can be established during transcriptional repression in yeast (Yadon et al., 2013) and murine cells (Jing et al., 2008). In the case of the former, it is thought that looping juxtaposes sites bound by the repressive chromatin remodeler Isw2 with silenced genes. In the case of the latter, acute repression of the Kit gene was associated not only with a loss of the enhancer–promoter loop but also with a concomitant gain of a repression-specific interaction of a promoter-proximal region with an intronic segment. Whether de novo loop formation at the Kit gene contributes to repression remains an open question. It is possible that alternative looped contacts compete with activating loops to diminish enhancer–promoter contacts. This indicates that loops are dynamic and that chromatin architectural changes seem as tightly coupled to gene repression as to activation.

Which proteins establish and maintain chromatin loops?

A longstanding and difficult to answer question relates to the identity of nuclear factors that forge chromatin loops. Early in vitro studies using electron microscopy showed that purified transcription factors capable of dimerization such as Sp1 or the viral protein E2 can form loops when bound to chromatin-free DNA templates (Knight et al., 1991; Su et al., 1991). Pinpointing the proteins at the base of the loops that form the actual connections in vivo remains an unresolved issue. Nonetheless, several proteins have been identified that are thought to contribute to chromatin looping, and a short summary of different factors that were found at the base of dynamic and/or stable long-range contacts will be provided. While some proteins act in a more cell type–specific manner (e.g., GATA1, Ldb1, and EKLF at the β-globin locus in erythroblasts [Drissen et al., 2004; Vakoc et al., 2005; Song et al., 2007], TAF3 in mESCs [Liu et al., 2011], and SATB1 in thymocytes [Cai et al., 2006]), others are thought to serve as more general looping factors (e.g., Mediator, CTCF, cohesin, and YY1 [ Rollins et al., 1999; Splinter et al., 2006; Kagey et al., 2010; Lai et al., 2013; Phillips-Cremins et al., 2013; Ing-Simmons et al., 2015; Beagan et al., 2017]).

ChIA-PET and HiChIP are methods that enrich for chromatin contacts associated with selected proteins or chromatin modifications of interest (Fullwood et al., 2009; Mumbach et al., 2016). These approaches have revealed that transcription factors (e.g., estrogen receptor; Fullwood et al., 2009), architectural proteins (e.g., CTCF and cohesin; Tang et al., 2015; Mumbach et al., 2016), and pol2 or histone acetylation (Zhang et al., 2013; Mumbach et al., 2017) can be linked to contacts among multiple enhancers and/or promoters in cis and in trans, thereby creating regulatory nodes, which led to the speculation that transcriptional regulation of multiple genes might be coordinated. However, the degree to which chromatin interactions between chromosomes impact gene expression remains a subject of debate (Cremer and Cremer, 2001; Meaburn and Misteli, 2007; Williams et al., 2010; Cavalli and Misteli, 2013; Bonev and Cavalli, 2016).

While the above reports provide insights into which factors and chromatin marks occupy the base of the loops, they do not establish evidence for their direct involvement as “glue” between contact sites. Studies that monitor in vivo long-range interactions have typically relied on loss-of-function assays of candidate looping factors or regulatory elements themselves (Drissen et al., 2004; Patrinos et al., 2004; Vakoc et al., 2005). However, these approaches typically fail to distinguish direct from secondary effects that might result from transcriptional perturbations. Even in the case of targeted tethering of the candidate looping factor Ldb1 to a predetermined site in the genome, which was successful in forging long-range chromatin contacts in the globin locus as discussed, definitive proof that this occurs in vivo via direct contacts rather than protein intermediates is difficult to attain. This challenge still stands for most, if not all, factors implicated in chromatin looping.

Enhancer function is restricted by insulators

Most enhancers are promiscous since they are capable of augmenting the expression of commonly used reporter genes with minimal promoters (Picard and Schaffner, 1983). Methods to identify enhancers rely on this concept (e.g., Arnold et al., 2013; Symmons et al., 2014). Moreover, as discussed, ectopic integration of the β-globin LCR into a novel genomic region leads to close association and activation of many but not all nearby genes, including nonerythroid ones (Noordermeer et al., 2008). What, then, underlies the high degree of specifity of enhancer action in vivo? One mode to constrain enhancer function comes from enhancer-blocking insulators (West et al., 2002). Insulators are position-dependent as their placement between an enhancer and promoter nullifies enhancer activity, while placement upstream or downstream is inoccuous (Fig. 3; West et al., 2002). This strict position dependence distinguishes them from transcriptional repressor elements (such as PREs) that can silence genes in a more position-independent manner. Interestingly, insulators were found to be able to reposition and colocalize PRE-repressed genes in Drosophila, thereby ensuring their silencing (Sigrist and Pirrotta, 1997; Comet et al., 2011; Li et al., 2011, 2013).

CTCF is currently thought to be the major enhancer blocking insulator protein in mammals (Bell et al., 1999). Thus, even though CTCF can play a role in fostering enhancer–promoter communication, it can also prevent it. This has been well characterized at the imprinting control region (ICR) between the paternally expressed Igf2 and the maternally expressed H19 genes, which operates in part via DNA methylation–sensitive CTCF binding (Bell and Felsenfeld, 2000; Hark et al., 2000; Engel et al., 2004). On the maternal chromosome, CTCF occupies the unmethylated ICR to shield the Igf2 gene from a downstream enhancer. On the paternal chromosome, the ICR is methylated, preventing CTCF binding and enabling the enhancer to interact with the Igf2 promoter (Murrell et al., 2004b; Kurukuti et al., 2006). In addition, it has been proposed that CTCF forms looped contacts with the blocked enhancer (Fig. 3; Yoon et al., 2007). Another instructive example of CTCF insulating an enhancer, presumably via loop formation, is provided by ectopic insertion of a CTCF-bound element between the β-globin genes and the LCR (Hou et al., 2008). The ectopic element pairs with a CTCF-bound element upstream of the LCR, encasing it in a loop (Fig. 3), which is thought to prevent LCR contacts with the globin promoters. Pairing of insulators has been described in Drosophila (Blanton et al., 2003; Byrd and Corces, 2003), but how widespread this dual function of CTCF of preventing loops while engaging in new loops is in mammals remains unclear. Loop formation via pairing of CTCF sites might not be required for enhancer-blocking activity, since it can occur in reporter assays with single CTCF elements and no known partner elements (Bell et al., 1999). CTCF might alternatively prevent tracking of an enhancer toward a promoter (Zhao and Dean, 2004), which would explain the position dependence of an enhancer blocker and at the same time provide a rationale as to why enhancers cannot simply loop across an insulator element to reach a promoter. This suggests that the role of CTCF as an insulator may involve looping in some but not all cases. Regardless, loss of CTCF can lead to aberrant gene expression, presumably due to enhancer–promoter miswiring.

Originally appreciated as a factor important for sister chromatid cohesion (Michaelis et al., 1997), cohesin was found to colocalize at >50% of CTCF sites across multiple cell lines (Parelho et al., 2008; Rubio et al., 2008; Wendt et al., 2008). It is likely to contribute to the insulation function of CTCF through mediating long-range chromatin interactions. Similarly, the insulating potential of CTCF is dependent on other cofactors (Ghirlando and Felsenfeld, 2016). In addition, insulation can also be obtained in an ostensibly CTCF-independent manner. For instance, the β-globin LCR can function promiscuously under experimental conditions, so why does it not activate fetal-type globin genes in adult erythroid cells and vice versa? Fine scale chromosomal contact maps in human erythroid cells identified a regulatory element that engages in developmental stage–specific long-range chromatin contacts in a manner that insulates the “wrong” type of globin genes from contacting the LCR (Huang et al., 2017). Deletion of this element (which does not contain any detectable CTCF-occupied sites) in adult cells leads to increased LCR contacts with fetal genes and reactivates them. How this element functions is currently not understood, but it provides an example of a developmental stage- and tissue-specific architectural element that influences enhancer–promoter wiring. This further suggests that different architectural proteins, some of which may be unknown, cooperate to set up a framework to promote or prevent contacts between regulatory elements.

Architectural constraints define regulatory domains

The introduction of Hi-C has enabled the mapping of chromatin interactions genome-wide (Lieberman-Aiden et al., 2009). Contact maps revealed that chromosomes are partitioned into roughly megabase-scale TADs (Dixon et al., 2012; Nora et al., 2012; Sexton et al., 2012). TADs are highly conserved between cell types and species (Dixon et al., 2012; Ho et al., 2014; Vietri Rudan et al., 2015). Accordingly, recent analysis of late primate evolution revealed that boundaries are depleted for evolutionary changes that disrupt their function (Fudenberg and Pollard, 2018 ,Preprint). The invariant nature of these domains suggests they represent a more general principle of chromosomal organization and have little influence on tissue-specific gene expression programs. This is consistent with the view that specific transcriptional profiles are mainly driven by unique long-range contacts between regulatory elements within TADs as well as compartmentalization of chromatin contacts between TADs (Dixon et al., 2015; Beagan et al., 2016, 2017; Bonev et al., 2017). Uniformity of TADs across tissues also suggests that gene expression cannot be the only determinant of TAD formation even though it likely contributes to it (Bonev et al., 2017). Of note, TADs were detected with high-resolution imaging as well as more recently developed genomic methods that are independent of proximity ligation, suggesting these findings are not reflective of technical biases (Beagrie et al., 2017; Nir et al., 2018 ,Preprint; Wang et al., 2016; Quinodoz et al., 2018; Szabo et al., 2018). Sub-TADs, which are nested within larger TADs are more variable between cell types (Phillips-Cremins et al., 2013; Rao et al., 2014). Communication between regulatory elements is generally restricted to such contact domains because interactions across TAD or sub-TAD boundaries are depleted. This was corroborated by insertion of a LacZ reporter-based enhancer trap at random sites in the mouse genome (Symmons et al., 2014). The reporter genes were mostly responsive to enhancers within the same TAD, indicating that TADs are not just physical structures but function to constrain enhancer influence. The other side of the coin is that TADs can also foster long-range enhancer–promoter contacts. Integration of the Shh limb enhancer at different sites within its TAD showed that the enhancer functions largely independent of distance (Symmons et al., 2016). However, when the TAD was disrupted, the enhancers seemed to only function at shorter distances. Thus, physical constraints imposed by TAD boundaries might not only prevent aberrant enhancer–promoter pairing but also facilitate long-range contacts between them, a concept reminiscent of the bystander effects due to forced chromatin looping as was discussed.

In line with its insulating and looping potential, CTCF was found to be enriched at TAD boundaries (Dixon et al., 2012; Nora et al., 2012; Phillips-Cremins et al., 2013). While insulator function often refers to blocking the communication between cis-regulatory elements (enhancer blocking insulators) or limiting the spreading of repressive chromatin (barrier insulators), boundary annotations of topological domains are often contact-based (usually measured by Hi-C). While the different functions of insulators can be separated (Recillas-Targa et al., 2002; West et al., 2004), they can be related, and as a result, the definition of these functions has become somewhat blurred. In general, chromatin interactions across insulator/boundary elements are disfavored, and CTCF is often found at the base of loops that define domains (loop domains; Rao et al., 2014). However, not all CTCF-occupied sites are boundaries (Dixon et al., 2012; Nora et al., 2012), which suggests that CTCF binding alone is insufficient for (sub)TAD boundary formation. CTCF binding per se can be tissue-specific, modulated by contextual transcription factors (e.g., Behera et al., 2018), but there are also examples of sites where CTCF is bound across a range of tissues but engaged in looped interactions only in one of them. This is the case, for instance, for CTCF sites flanking the α- and β-globin loci (e.g., Hou et al., 2008; Hanssen et al., 2017; Huang et al., 2017).

The functional relevance of domain boundaries in safeguarding proper gene regulation has been impressively demonstrated in cases where their disruption leads to disease (reviewed in Krijger and de Laat, 2016). Structural variants, such as inversion, duplication, or deletion of a CTCF-associated TAD boundary can result in pairing between a limb enhancer and a gene normally not regulated by this enhancer, leading to digit malformation (Lupiáñez et al., 2015). Cook’s syndrome, characterized by limb malformations, can be caused by a specific duplication event that forms a new TAD in which the KCNJ2 gene is contacted and activated by SOX9 enhancers (Franke et al., 2016). Targeted deletion of CTCF sites at boundaries near the miR-290-295 and Pou5f1 loci in mESCs enabled new enhancer contacts and activation of nearby genes (Dowen et al., 2014). Similar observations have been made at a boundary near an erythroid gene where deletion of a CTCF site enables a tissue- and developmental stage–specific enhancer to act on a housekeeping gene (Hsu et al., 2017). At the HoxA-locus, deletion of a CTCF site leads to spreading of active chromatin into a previously repressed domain and inappropriate gene activation (Narendra et al., 2015). In patients with T cell acute lymphoblastic leukemia, microdeletions perturb a CTCF-associated boundary, which results in proto-oncogene activation (Hnisz et al., 2016). In gain-of-function IDH mutant gliomas, DNA hypermethylation prevents CTCF binding and perturbs a domain boundary, allowing an enhancer to aberrantly activate a proto-oncogene (Flavahan et al., 2016). Deletion of a CTCF–cohesin cobound site that demarcates one end of an erythroid-specific sub-TAD surrounding the mouse α-globin locus extended the sub-TAD, enabling an α-globin regulatory element to contact and augment the expression of genes outside the sub-TAD (Hanssen et al., 2017). These examples illustrate how CTCF loss can lead to enhancer–promoter miswiring.

It may, however, be over-simplified to conceptually equate domain boundaries with functional insulation by CTCF. Numerous recent papers show that features such as housekeeping genes, pol2 occupancy, transcriptional activity, and short interspersed nuclear elements are also enriched at TAD boundaries (Dixon et al., 2012; Nora et al., 2012; Rowley et al., 2017). Transposable element-mediated expansion of transcription factor binding sites (such as for CTCF) was found to be a substrate for cooption of regulatory functions during evolution. Therefore, CTCF and gene activity may (co)evolve to establish regulatory domains (Schmidt et al., 2012; Sundaram et al., 2014).

Global perturbation of architectural proteins and the 3D genome

The previously described examples at single loci highlight the potential consequences of perturbing (CTCF) boundaries, but recent experiments in which architectural proteins were depleted globally report limited transcriptional changes, even though domain configuration was markedly perturbed. Auxin-mediated degradation of CTCF for 24 h in mESCs decreased domain insulation at around 80% of the 5,525 boundaries that were called, but only ∼400 genes were transcriptionally affected (Nora et al., 2017). Approximately half of the genes whose expression changed were down-regulated. Those tended to have CTCF bound close to their transcriptional start site. Next to loss of CTCF as a potential transcriptional activator (West et al., 2002), expression changes may result from misregulation of CTCF-mediated loops that arrange pairing among cis-regulatory elements (Phillips and Corces, 2009; Ghirlando and Felsenfeld, 2016). The genes that were up-regulated upon CTCF depletion were often located close to boundaries, raising the possibility that they came under the inappropriate influence of enhancers. However, gene activation was rare under these conditions (Nora et al., 2017), suggesting that other requirements, such as the presence of certain transcription factors, may have to be met in specific cases in order for aberrant transcription to occur even when genome partitioning defects spatially allow for it. This is the case for the β-globin locus, for example, where mutations or deletion of CTCF sites led to new interactions, but did not result in activation of surrounding nonerythroid genes by the LCR (Bulger et al., 2003; Splinter et al., 2006).

Contact domains have been proposed to be formed by a mechanism called loop extrusion (Nasmyth, 2001; Sanborn et al., 2015; Fudenberg et al., 2016). Upon recruitment to genomic elements, the cohesin complexes are thought to propel the chromatin fiber through their ring-shaped structure until extrusion stalls at convergently oriented CTCF sites (de Wit et al., 2015; Guo et al., 2015; Vietri Rudan et al., 2015). This, in concert with tightly regulated loading and unloading of extrusion factors, results in looped domain-like configurations. This model found support in studies in which cohesin subunits and the factors that load and unload components of the complex were targeted. Knockout of the cohesin unloading factor WAPL1 in HAP1 cells increased the size of loops between convergent CTCF sites due to loop extrusion beyond primary CTCF sites at which it would normally stall (Haarhuis et al., 2017). Conversely, knockout of the cohesin subunit Rad21 or its loading factor NIPBL resulted in loss of TADs and revealed a 3D organization that was more reflective of compartments based on epigenomic chromatin landscapes (Rao et al., 2017; Schwarzer et al., 2017). This is in line with recent findings in Drosophila, in which genes with comparable transcriptional status were found to converge in mini-domains (Rowley et al., 2017). In-depth analysis of Hi-C data in a human lymphoblastoid cell line revealed comparable active/inactive switches at domain borders. With CTCF present at the majority of these borders (Rowley et al., 2017), this corroborates the interplay between architectural proteins and transcriptional activity in domain formation.

Early embryonic lethality of CTCF knockout mice suggests that this protein is indispensable for proper gene regulation (Moore et al., 2012). In addition, cell death is observed in mammalian cells upon prolonged CTCF depletion, and cohesin loss interferes with cell division (Soshnikova et al., 2010; Watson et al., 2014; Gupta et al., 2016; Nora et al., 2017). However, CTCF is not required for embryonic development in Drosophila (Gambetta and Furlong, 2018), and while pronounced developmental and disease phenotypes at the organismal level are observed in individuals with mutations in genes that code for cohesin subunits, these alterations do not prevent viability (Watrin et al., 2016). As discussed, acute depletion of architectural proteins impaired 3D folding without dramatically altering the transcriptome. The ostensibly mild changes at the level of gene expression after domain disruptions raise questions regarding the degree to which large-scale chromatin architecture impinges on gene expression. One needs to keep in mind, however, that boundary loss has been measured mainly by Hi-C. As suggested by recent super-resolution imaging (Bintu et al., 2018), it is possible that smaller shifts and heterogeneity at boundaries as a result of architectural protein depletion might lead to loss of boundary calls in cell population–based experiments even though boundaries could be functionally preserved at the single-cell level. Together with the other examples in which boundary loss does lead to enhancer–promoter miswiring and disease, this suggests that consequences may have to be investigated on a case-by-case basis because of context specificity.

Conclusion

In this review, we attempted to highlight examples of the closely intertwined relationship between chromosomal architectural features and gene regulation. While most studies, especially earlier ones, are correlative when linking gene activity to nuclear architecture, an increasing number includes specifically targeted perturbations, allowing inferences about causal relationships. We have learned that gene positioning, chromosomal looping, and gene expression are in mutually influential relationships. However, the state of a gene cannot predict architectural features surrounding it with certainty and vice versa. For example, as illustrated above, escape from the nuclear periphery, extrusion from the chromosome territory, and enhancer looping are frequently seen at active genes, but none of these features are fully deterministic. Moreover, the interplay between regulatory elements and their nuclear environment may be distinct between genes, where, for example, in one scenario, enhancer–promoter loops exist before activation, whereas in another, looping might be rate-limiting. These considerations are also important when using chromosomal contact maps to interpret genome-wide association study signals and making inferences of nucleotide variants on variable traits or disease mechanisms.

While this field has witnessed a massive expansion in knowledge in a relatively short time span, key challenges remain. These include establishing the identity of nuclear factors that directly assemble the myriad of chromosomal configurations observed. This issue is not easily addressed since depletion, even transiently, of nuclear factors can have wide-ranging secondary effects, and since even precise editing of transcription factor binding sites can affect chromatin association of nearby factors. Hence, loss-of-function and gain-of-function studies have to be combined to strengthen the conclusions. Another important issue that the field has to contend with is that while general principles of chromatin organization and gene regulation have been recognized, there are clearly differences in the way different regulatory elements function and genes are controlled. This necessitates deep reductionist experimentation at individual loci if the goal is to fully understand a specific gene.

Another widely faced challenge in the field is cell-to-cell and even allele-to-allele variability in gene regulation and architecture. Heterogeneity in gene expression is impinged upon by responsiveness to regulatory cues (e.g., signaling gradients, asymmetric distribution of cellular content during mitosis, etc.), cell cycle stage, and stochastic effects. Especially when it comes to the cell cycle, everything that is measurable about higher-order nuclear architecture undergoes dramatic reorganization during the life of a cell, most strikingly during mitosis. This needs to be taken into account when targeted experimental perturbations can affect cell cycle progression.

The road ahead promises that many of the above challenges can be met. This optimisim is rooted in the stunning improvements of single-cell technologies and live imaging tools, as well as more refined genome and epigenome editing methods. Hence, our insights into the intricacies of the nucleus will benefit from ever finer spatial and temporal resolution.

Acknowledgments

We thank Jennifer E. Phillips-Cremins, Peter H.L. Krijger, and Haoyue Zhang for critically reading the manuscript and for helpful suggestions.

Cited work from the laboratory of G.A. Blobel is supported by grants from the National Institutes of Health (RO1DK54937, R01HL119479, R24DK106766, U01HL129998A2017, and R37DK058044). M.W. Vermunt is supported by the Rubicon research program, which is financed by the Netherlands Organization for Scientific Research (project no. 019.173EN.006).

The authors declare no competing financial interests.

References

References
Aguilar-Arnal
,
L.
,
O.
Hakim
,
V.R.
Patel
,
P.
Baldi
,
G.L.
Hager
, and
P.
Sassone-Corsi
.
2013
.
Cycles in spatial and temporal chromosomal organization driven by the circadian clock
.
Nat. Struct. Mol. Biol.
20
:
1206
1213
.
Akhtar
,
A.
, and
S.M.
Gasser
.
2007
.
The nuclear envelope and transcriptional control
.
Nat. Rev. Genet.
8
:
507
517
.
Amano
,
T.
,
T.
Sagai
,
H.
Tanabe
,
Y.
Mizushina
,
H.
Nakazawa
, and
T.
Shiroishi
.
2009
.
Chromosomal dynamics at the Shh locus: limb bud-specific differential regulation of competence and active transcription
.
Dev. Cell.
16
:
47
57
.
Andrey
,
G.
,
R.
Schöpflin
,
I.
Jerković
,
V.
Heinrich
,
D.M.
Ibrahim
,
C.
Paliou
,
M.
Hochradel
,
B.
Timmermann
,
S.
Haas
,
M.
Vingron
, and
S.
Mundlos
.
2017
.
Characterization of hundreds of regulatory landscapes in developing limbs reveals two regimes of chromatin folding
.
Genome Res.
27
:
223
233
.
Andrulis
,
E.D.
,
A.M.
Neiman
,
D.C.
Zappulla
, and
R.
Sternglanz
.
1998
.
Perinuclear localization of chromatin facilitates transcriptional silencing
.
Nature.
394
:
592
595
.
Arnold
,
C.D.
,
D.
Gerlach
,
C.
Stelzer
,
Ł.M.
Boryń
,
M.
Rath
, and
A.
Stark
.
2013
.
Genome-wide quantitative enhancer activity maps identified by STARR-seq
.
Science.
339
:
1074
1077
.
Banerji
,
J.
,
S.
Rusconi
, and
W.
Schaffner
.
1981
.
Expression of a beta-globin gene is enhanced by remote SV40 DNA sequences
.
Cell.
27
:
299
308
.
Bartman
,
C.R.
, and
G.A.
Blobel
.
2015
.
Perturbing Chromatin Structure to Understand Mechanisms of Gene Expression
.
Cold Spring Harb. Symp. Quant. Biol.
80
:
207
212
.
Bartman
,
C.R.
,
S.C.
Hsu
,
C.C.-S.
Hsiung
,
A.
Raj
, and
G.A.
Blobel
.
2016
.
Enhancer Regulation of Transcriptional Bursting Parameters Revealed by Forced Chromatin Looping
.
Mol. Cell.
62
:
237
247
.
Beagan
,
J.A.
,
T.G.
Gilgenast
,
J.
Kim
,
Z.
Plona
,
H.K.
Norton
,
G.
Hu
,
S.C.
Hsu
,
E.J.
Shields
,
X.
Lyu
,
E.
Apostolou
, et al
2016
.
Local Genome Topology Can Exhibit an Incompletely Rewired 3D-Folding State during Somatic Cell Reprogramming
.
Cell Stem Cell.
18
:
611
624
.
Beagan
,
J.A.
,
M.T.
Duong
,
K.R.
Titus
,
L.
Zhou
,
Z.
Cao
,
J.
Ma
,
C.V.
Lachanski
,
D.R.
Gillis
, and
J.E.
Phillips-Cremins
.
2017
.
YY1 and CTCF orchestrate a 3D chromatin looping switch during early neural lineage commitment
.
Genome Res.
27
:
1139
1152
.
Beagrie
,
R.A.
,
A.
Scialdone
,
M.
Schueler
,
D.C.A.
Kraemer
,
M.
Chotalia
,
S.Q.
Xie
,
M.
Barbieri
,
I.
de Santiago
,
L.-M.
Lavitas
,
M.R.
Branco
, et al
2017
.
Complex multi-enhancer contacts captured by genome architecture mapping
.
Nature.
543
:
519
524
.
Behera
,
V.
,
P.
Evans
,
C.J.
Face
,
N.
Hamagami
,
L.
Sankaranarayanan
,
C.A.
Keller
,
B.
Giardine
,
K.
Tan
,
R.C.
Hardison
,
J.
Shi
, and
G.A.
Blobel
.
2018
.
Exploiting genetic variation to uncover rules of transcription factor binding and chromatin accessibility
.
Nat. Commun.
9
:
782
.
Bell
,
A.C.
, and
G.
Felsenfeld
.
2000
.
Methylation of a CTCF-dependent boundary controls imprinted expression of the Igf2 gene
.
Nature.
405
:
482
485
.
Bell
,
A.C.
,
A.G.
West
, and
G.
Felsenfeld
.
1999
.
The protein CTCF is required for the enhancer blocking activity of vertebrate insulators
.
Cell.
98
:
387
396
.
Benabdallah
,
N.S.
,
I.
Williamson
,
R.S.
Illingworth
,
S.
Boyle
,
G.R.
Grimes
,
P.
Therizols
, and
W.
Bickmore
.
2017
.
PARP mediated chromatin unfolding is coupled to long-range enhancer activation
.
bioRxiv.
doi: (Preprint posted June 25, 2017)
Bender
,
M.A.
,
A.
Reik
,
J.
Close
,
A.
Telling
,
E.
Epner
,
S.
Fiering
,
R.
Hardison
, and
M.
Groudine
.
1998
.
Description and targeted deletion of 5′ hypersensitive site 5 and 6 of the mouse beta-globin locus control region
.
Blood.
92
:
4394
4403
.
Bender
,
M.A.
,
J.N.
Roach
,
J.
Halow
,
J.
Close
,
R.
Alami
,
E.E.
Bouhassira
,
M.
Groudine
, and
S.N.
Fiering
.
2001
.
Targeted deletion of 5‘HS1 and 5’HS4 of the beta-globin locus control region reveals additive activity of the DNaseI hypersensitive sites
.
Blood.
98
:
2022
2027
.
Berrozpe
,
G.
,
G.O.
Bryant
,
K.
Warpinski
,
D.
Spagna
,
S.
Narayan
,
S.
Shah
, and
M.
Ptashne
.
2017
.
Polycomb Responds to Low Levels of Transcription
.
Cell Reports.
20
:
785
793
.
Bickmore
,
W.A.
, and
B.
van Steensel
.
2013
.
Genome architecture: domain organization of interphase chromosomes
.
Cell.
152
:
1270
1284
.
Bintu
,
B.
,
L.J.
Mateo
,
J.-H.
Su
,
N.A.
Sinnott-Armstrong
,
M.
Parker
,
S.
Kinrot
,
K.
Yamaya
,
A.N.
Boettiger
, and
X.
Zhuang
.
2018
.
Super-resolution chromatin tracing reveals domains and cooperative interactions in single cells
.
Science.
362
:
eaau11783
.
Blanton
,
J.
,
M.
Gaszner
, and
P.
Schedl
.
2003
.
Protein:protein interactions and the pairing of boundary elements in vivo
.
Genes Dev.
17
:
664
675
.
Boettiger
,
A.N.
,
B.
Bintu
,
J.R.
Moffitt
,
S.
Wang
,
B.J.
Beliveau
,
G.
Fudenberg
,
M.
Imakaev
,
L.A.
Mirny
,
C.-T.
Wu
, and
X.
Zhuang
.
2016
.
Super-resolution imaging reveals distinct chromatin folding for different epigenetic states
.
Nature.
529
:
418
422
.
Bonev
,
B.
, and
G.
Cavalli
.
2016
.
Organization and function of the 3D genome
.
Nat. Rev. Genet.
17
:
772
772
.
Bonev
,
B.
,
N.
Mendelson Cohen
,
Q.
Szabo
,
L.
Fritsch
,
G.L.
Papadopoulos
,
Y.
Lubling
,
X.
Xu
,
X.
Lv
,
J.-P.
Hugnot
,
A.
Tanay
, and
G.
Cavalli
.
2017
.
Multiscale 3D Genome Rewiring during Mouse Neural Development
.
Cell.
171
:
557
572.e24
.
Buecker
,
C.
, and
J.
Wysocka
.
2012
.
Enhancers as information integration hubs in development: lessons from genomics
.
Trends Genet.
28
:
276
284
.
Bulger
,
M.
, and
M.
Groudine
.
2011
.
Functional and mechanistic diversity of distal transcription enhancers
.
Cell.
144
:
327
339
.
Bulger
,
M.
,
D.
Schübeler
,
M.A.
Bender
,
J.
Hamilton
,
C.M.
Farrell
,
R.C.
Hardison
, and
M.
Groudine
.
2003
.
A complex chromatin landscape revealed by patterns of nuclease sensitivity and histone modification within the mouse beta-globin locus
.
Mol. Cell. Biol.
23
:
5234
5244
.
Byrd
,
K.
, and
V.G.
Corces
.
2003
.
Visualization of chromatin domains created by the gypsy insulator of Drosophila
.
J. Cell Biol.
162
:
565
574
.
Cai
,
S.
,
C.C.
Lee
, and
T.
Kohwi-Shigematsu
.
2006
.
SATB1 packages densely looped, transcriptionally active chromatin for coordinated expression of cytokine genes
.
Nat. Genet.
38
:
1278
1288
.
Cajiao
,
I.
,
A.
Zhang
,
E.J.
Yoo
,
N.E.
Cooke
, and
S.A.
Liebhaber
.
2004
.
Bystander gene activation by a locus control region
.
EMBO J.
23
:
3854
3863
.
Calo
,
E.
, and
J.
Wysocka
.
2013
.
Modification of enhancer chromatin: what, how, and why?
Mol. Cell.
49
:
825
837
.
Carter
,
D.
,
L.
Chakalova
,
C.S.
Osborne
,
Y.-F.
Dai
, and
P.
Fraser
.
2002
.
Long-range chromatin regulatory interactions in vivo
.
Nat. Genet.
32
:
623
626
.
Cavalli
,
G.
, and
T.
Misteli
.
2013
.
Functional implications of genome topology
.
Nat. Struct. Mol. Biol.
20
:
290
299
.
Chambeyron
,
S.
, and
W.A.
Bickmore
.
2004
.
Chromatin decondensation and nuclear reorganization of the HoxB locus upon induction of transcription
.
Genes Dev.
18
:
1119
1130
.
Chan
,
C.S.
,
L.
Rastelli
, and
V.
Pirrotta
.
1994
.
A Polycomb response element in the Ubx gene that determines an epigenetically inherited state of repression
.
EMBO J.
13
:
2553
2564
.
Chen
,
H.
,
M.
Levo
,
L.
Barinov
,
M.
Fujioka
,
J.B.
Jaynes
, and
T.
Gregor
.
2018
.
Dynamic interplay between enhancer-promoter topology and gene activity
.
Nat. Genet.
50
:
1296
1303
.
Cho
,
S.W.
,
J.
Xu
,
R.
Sun
,
M.R.
Mumbach
,
A.C.
Carter
,
Y.G.
Chen
,
K.E.
Yost
,
J.
Kim
,
J.
He
,
S.A.
Nevins
, et al
2018
.
Promoter of lncRNA Gene PVT1 Is a Tumor-Suppressor DNA Boundary Element
.
Cell.
173
:
1398
1412.e22
.
Chopra
,
V.S.
,
N.
Kong
, and
M.
Levine
.
2012
.
Transcriptional repression via antilooping in the Drosophila embryo
.
Proc. Natl. Acad. Sci. USA.
109
:
9460
9464
.
Comet
,
I.
,
B.
Schuettengruber
,
T.
Sexton
, and
G.
Cavalli
.
2011
.
A chromatin insulator driving three-dimensional Polycomb response element (PRE) contacts and Polycomb association with the chromatin fiber
.
Proc. Natl. Acad. Sci. USA.
108
:
2294
2299
.
Cremer
,
T.
, and
C.
Cremer
.
2001
.
Chromosome territories, nuclear architecture and gene regulation in mammalian cells
.
Nat. Rev. Genet.
2
:
292
301
.
Cremer
,
T.
,
M.
Cremer
,
S.
Dietzel
,
S.
Müller
,
I.
Solovei
, and
S.
Fakan
.
2006
.
Chromosome territories--a functional nuclear landscape
.
Curr. Opin. Cell Biol.
18
:
307
316
.
Cullen
,
K.E.
,
M.P.
Kladde
, and
M.A.
Seyfred
.
1993
.
Interaction between transcription regulatory regions of prolactin chromatin
.
Science.
261
:
203
206
.
De Gobbi
,
M.
,
V.
Viprakasit
,
J.R.
Hughes
,
C.
Fisher
,
V.J.
Buckle
,
H.
Ayyub
,
R.J.
Gibbons
,
D.
Vernimmen
,
Y.
Yoshinaga
,
P.
de Jong
, et al
2006
.
A regulatory SNP causes a human genetic disease by creating a new transcriptional promoter
.
Science.
312
:
1215
1217
.
Dekker
,
J.
,
K.
Rippe
,
M.
Dekker
, and
N.
Kleckner
.
2002
.
Capturing chromosome conformation
.
Science.
295
:
1306
1311
.
de Laat
,
W.
, and
D.
Duboule
.
2013
.
Topology of mammalian developmental enhancers and their regulatory landscapes
.
Nature.
502
:
499
506
.
Deng
,
W.
, and
G.A.
Blobel
.
2014
.
Manipulating nuclear architecture
.
Curr. Opin. Genet. Dev.
25
:
1
7
.
Deng
,
W.
,
J.
Lee
,
H.
Wang
,
J.
Miller
,
A.
Reik
,
P.D.
Gregory
,
A.
Dean
, and
G.A.
Blobel
.
2012
.
Controlling long-range genomic interactions at a native locus by targeted tethering of a looping factor
.
Cell.
149
:
1233
1244
.
Deng
,
W.
,
J.W.
Rupon
,
I.
Krivega
,
L.
Breda
,
I.
Motta
,
K.S.
Jahn
,
A.
Reik
,
P.D.
Gregory
,
S.
Rivella
,
A.
Dean
, and
G.A.
Blobel
.
2014
.
Reactivation of developmentally silenced globin genes by forced chromatin looping
.
Cell.
158
:
849
860
.
Denker
,
A.
, and
W.
de Laat
.
2016
.
The second decade of 3C technologies: detailed insights into nuclear organization
.
Genes Dev.
30
:
1357
1382
.
de Wit
,
E.
,
E.S.M.
Vos
,
S.J.B.
Holwerda
,
C.
Valdes-Quezada
,
M.J.A.M.
Verstegen
,
H.
Teunissen
,
E.
Splinter
,
P.J.
Wijchers
,
P.H.L.
Krijger
, and
W.
de Laat
.
2015
.
CTCF Binding Polarity Determines Chromatin Looping
.
Mol. Cell.
60
:
676
684
.
Dixon
,
J.R.
,
S.
Selvaraj
,
F.
Yue
,
A.
Kim
,
Y.
Li
,
Y.
Shen
,
M.
Hu
,
J.S.
Liu
, and
B.
Ren
.
2012
.
Topological domains in mammalian genomes identified by analysis of chromatin interactions
.
Nature.
485
:
376
380
.
Dixon
,
J.R.
,
I.
Jung
,
S.
Selvaraj
,
Y.
Shen
,
J.E.
Antosiewicz-Bourget
,
A.Y.
Lee
,
Z.
Ye
,
A.
Kim
,
N.
Rajagopal
,
W.
Xie
, et al
2015
.
Chromatin architecture reorganization during stem cell differentiation
.
Nature.
518
:
331
336
.
Dowen
,
J.M.
,
Z.P.
Fan
,
D.
Hnisz
,
G.
Ren
,
B.J.
Abraham
,
L.N.
Zhang
,
A.S.
Weintraub
,
J.
Schujiers
,
T.I.
Lee
,
K.
Zhao
, and
R.A.
Young
.
2014
.
Control of cell identity genes occurs in insulated neighborhoods in mammalian chromosomes
.
Cell.
159
:
374
387
.
Drissen
,
R.
,
R.-J.
Palstra
,
N.
Gillemans
,
E.
Splinter
,
F.
Grosveld
,
S.
Philipsen
, and
W.
de Laat
.
2004
.
The active spatial organization of the beta-globin locus requires the transcription factor EKLF
.
Genes Dev.
18
:
2485
2490
.
Duncan
,
I.M.
1982
.
Polycomblike: a gene that appears to be required for the normal expression of the bithorax and antennapedia gene complexes of Drosophila melanogaster
.
Genetics.
102
:
49
70
.
Engel
,
N.
,
A.G.
West
,
G.
Felsenfeld
, and
M.S.
Bartolomei
.
2004
.
Antagonism between DNA hypermethylation and enhancer-blocking activity at the H19 DMD is uncovered by CpG mutations
.
Nat. Genet.
36
:
883
888
.
Engreitz
,
J.M.
,
A.
Pandya-Jones
,
P.
McDonel
,
A.
Shishkin
,
K.
Sirokman
,
C.
Surka
,
S.
Kadri
,
J.
Xing
,
A.
Goren
,
E.S.
Lander
, et al
2013
.
The Xist lncRNA exploits three-dimensional genome architecture to spread across the X chromosome
.
Science.
341
:
1237973
.
Fang
,
X.
,
J.
Sun
,
P.
Xiang
,
M.
Yu
,
P.A.
Navas
,
K.R.
Peterson
,
G.
Stamatoyannopoulos
, and
Q.
Li
.
2005
.
Synergistic and additive properties of the beta-globin locus control region (LCR) revealed by 5'HS3 deletion mutations: implication for LCR chromatin architecture
.
Mol. Cell. Biol.
25
:
7033
7041
.
Finlan
,
L.E.
,
D.
Sproul
,
I.
Thomson
,
S.
Boyle
,
E.
Kerr
,
P.
Perry
,
B.
Ylstra
,
J.R.
Chubb
, and
W.A.
Bickmore
.
2008
.
Recruitment to the nuclear periphery can alter expression of genes in human cells
.
PLoS Genet.
4
:
e1000039
.
Flavahan
,
W.A.
,
Y.
Drier
,
B.B.
Liau
,
S.M.
Gillespie
,
A.S.
Venteicher
,
A.O.
Stemmer-Rachamimov
,
M.L.
Suvà
, and
B.E.
Bernstein
.
2016
.
Insulator dysfunction and oncogene activation in IDH mutant gliomas
.
Nature.
529
:
110
114
.
Foley
,
K.P.
, and
J.D.
Engel
.
1992
.
Individual stage selector element mutations lead to reciprocal changes in beta- vs. epsilon-globin gene transcription: genetic confirmation of promoter competition during globin gene switching
.
Genes Dev.
6
:
730
744
.
Francastel
,
C.
,
M.C.
Walters
,
M.
Groudine
, and
D.I.
Martin
.
1999
.
A functional enhancer suppresses silencing of a transgene and prevents its localization close to centrometric heterochromatin
.
Cell.
99
:
259
269
.
Franke
,
M.
,
D.M.
Ibrahim
,
G.
Andrey
,
W.
Schwarzer
,
V.
Heinrich
,
R.
Schöpflin
,
K.
Kraft
,
R.
Kempfer
,
I.
Jerković
,
W.-L.
Chan
, et al
2016
.
Formation of new chromatin domains determines pathogenicity of genomic duplications
.
Nature.
538
:
265
269
.
Freire-Pritchett
,
P.
,
S.
Schoenfelder
,
C.
Várnai
,
S.W.
Wingett
,
J.
Cairns
,
A.J.
Collier
,
R.
García-Vílchez
,
M.
Furlan-Magaril
,
C.S.
Osborne
,
P.
Fraser
, et al
2017
.
Global reorganisation of cis-regulatory units upon lineage commitment of human embryonic stem cells
.
eLife.
6
:
699
.
Fudenberg
,
G.
, and
K.
Pollard
.
2018
.
Chromatin features constrain structural variation across evolutionary timescales
.
bioRxiv.
(Preprint posted April 30, 2018)
Fudenberg
,
G.
,
M.
Imakaev
,
C.
Lu
,
A.
Goloborodko
,
N.
Abdennur
, and
L.A.
Mirny
.
2016
.
Formation of Chromosomal Domains by Loop Extrusion
.
Cell Reports.
15
:
2038
2049
.
Fukaya
,
T.
,
B.
Lim
, and
M.
Levine
.
2016
.
Enhancer Control of Transcriptional Bursting
.
Cell.
166
:
358
368
.
Fulco
,
C.P.
,
M.
Munschauer
,
R.
Anyoha
,
G.
Munson
,
S.R.
Grossman
,
E.M.
Perez
,
M.
Kane
,
B.
Cleary
,
E.S.
Lander
, and
J.M.
Engreitz
.
2016
.
Systematic mapping of functional enhancer-promoter connections with CRISPR interference
.
Science.
354
:
769
773
.
Fullwood
,
M.J.
,
M.H.
Liu
,
Y.F.
Pan
,
J.
Liu
,
H.
Xu
,
Y.B.
Mohamed
,
Y.L.
Orlov
,
S.
Velkov
,
A.
Ho
,
P.H.
Mei
, et al
2009
.
An oestrogen-receptor-alpha-bound human chromatin interactome
.
Nature.
462
:
58
64
.
Gambetta
,
M.C.
, and
E.E.M.
Furlong
.
2018
.
The Insulator Protein CTCF Is Required for Correct Hox Gene Expression, but Not for Embryonic Development in Drosophila
.
Genetics.
210
:
129
136
.
Ghirlando
,
R.
, and
G.
Felsenfeld
.
2016
.
CTCF: making the right connections
.
Genes Dev.
30
:
881
891
.
Guo
,
Y.
,
Q.
Xu
,
D.
Canzio
,
J.
Shou
,
J.
Li
,
D.U.
Gorkin
,
I.
Jung
,
H.
Wu
,
Y.
Zhai
,
Y.
Tang
, et al
2015
.
CRISPR Inversion of CTCF Sites Alters Genome Topology and Enhancer/Promoter Function
.
Cell.
162
:
900
910
.
Gupta
,
P.
,
T.
Lavagnolli
,
H.
Mira-Bontenbal
,
A.G.
Fisher
, and
M.
Merkenschlager
.
2016
.
Cohesin’s role in pluripotency and reprogramming
.
Cell Cycle.
15
:
324
330
.
Haarhuis
,
J.H.I.
,
R.H.
van der Weide
,
V.A.
Blomen
,
J.O.
Yáñez-Cuna
,
M.
Amendola
,
M.S.
van Ruiten
,
P.H.L.
Krijger
,
H.
Teunissen
,
R.H.
Medema
,
B.
van Steensel
, et al
2017
.
The Cohesin Release Factor WAPL Restricts Chromatin Loop Extension
.
Cell.
169
:
693
707.e14
.
Hanssen
,
L.L.P.
,
M.T.
Kassouf
,
A.M.
Oudelaar
,
D.
Biggs
,
C.
Preece
,
D.J.
Downes
,
M.
Gosden
,
J.A.
Sharpe
,
J.A.
Sloane-Stanley
,
J.R.
Hughes
, et al
2017
.
Tissue-specific CTCF-cohesin-mediated chromatin architecture delimits enhancer interactions and function in vivo
.
Nat. Cell Biol.
19
:
952
961
.
Hark
,
A.T.
,
C.J.
Schoenherr
,
D.J.
Katz
,
R.S.
Ingram
,
J.M.
Levorse
, and
S.M.
Tilghman
.
2000
.
CTCF mediates methylation-sensitive enhancer-blocking activity at the H19/Igf2 locus
.
Nature.
405
:
486
489
.
Harr
,
J.C.
,
T.R.
Luperchio
,
X.
Wong
,
E.
Cohen
,
S.J.
Wheelan
, and
K.L.
Reddy
.
2015
.
Directed targeting of chromatin to the nuclear lamina is mediated by chromatin state and A-type lamins
.
J. Cell Biol.
208
:
33
52
.
Heinz
,
S.
,
C.E.
Romanoski
,
C.
Benner
, and
C.K.
Glass
.
2015
.
The selection and function of cell type-specific enhancers
.
Nat. Rev. Mol. Cell Biol.
16
:
144
154
.
Hnisz
,
D.
,
A.S.
Weintraub
,
D.S.
Day
,
A.-L.
Valton
,
R.O.
Bak
,
C.H.
Li
,
J.
Goldmann
,
B.R.
Lajoie
,
Z.P.
Fan
,
A.A.
Sigova
, et al
2016
.
Activation of proto-oncogenes by disruption of chromosome neighborhoods
.
Science.
351
:
1454
1458
.
Ho
,
J.W.K.
,
Y.L.
Jung
,
T.
Liu
,
B.H.
Alver
,
S.
Lee
,
K.
Ikegami
,
K.-A.
Sohn
,
A.
Minoda
,
M.Y.
Tolstorukov
,
A.
Appert
, et al
2014
.
Comparative analysis of metazoan chromatin organization
.
Nature.
512
:
449
452
.
Hou
,
C.
,
H.
Zhao
,
K.
Tanimoto
, and
A.
Dean
.
2008
.
CTCF-dependent enhancer-blocking by alternative chromatin loop formation
.
Proc. Natl. Acad. Sci. USA.
105
:
20398
20403
.
Hsu
,
S.C.
,
T.G.
Gilgenast
,
C.R.
Bartman
,
C.R.
Edwards
,
A.J.
Stonestrom
,
P.
Huang
,
D.J.
Emerson
,
P.
Evans
,
M.T.
Werner
,
C.A.
Keller
, et al
2017
.
The BET Protein BRD2 Cooperates with CTCF to Enforce Transcriptional and Architectural Boundaries
.
Mol. Cell.
66
:
102
116.e7
.
Huang
,
P.
,
C.A.
Keller
,
B.
Giardine
,
J.D.
Grevet
,
J.O.J.
Davies
,
J.R.
Hughes
,
R.
Kurita
,
Y.
Nakamura
,
R.C.
Hardison
, and
G.A.
Blobel
.
2017
.
Comparative analysis of three-dimensional chromosomal architecture identifies a novel fetal hemoglobin regulatory element
.
Genes Dev.
31
:
1704
1713
.
Ingham
,
P.W.
1985
.
Genetic control of the spatial pattern of selector gene expression in Drosophila
.
Cold Spring Harb. Symp. Quant. Biol.
50
:
201
208
.
Ing-Simmons
,
E.
,
V.C.
Seitan
,
A.J.
Faure
,
P.
Flicek
,
T.
Carroll
,
J.
Dekker
,
A.G.
Fisher
,
B.
Lenhard
, and
M.
Merkenschlager
.
2015
.
Spatial enhancer clustering and regulation of enhancer-proximal genes by cohesin
.
Genome Res.
25
:
504
513
.
Javierre
,
B.M.
,
O.S.
Burren
,
S.P.
Wilder
,
R.
Kreuzhuber
,
S.M.
Hill
,
S.
Sewitz
,
J.
Cairns
,
S.W.
Wingett
,
C.
Várnai
,
M.J.
Thiecke
, et al
BLUEPRINT Consortium
.
2016
.
Lineage-Specific Genome Architecture Links Enhancers and Non-coding Disease Variants to Target Gene Promoters
.
Cell.
167
:
1369
1384.e19
.
Jing
,
H.
,
C.R.
Vakoc
,
L.
Ying
,
S.
Mandat
,
H.
Wang
,
X.
Zheng
, and
G.A.
Blobel
.
2008
.
Exchange of GATA factors mediates transitions in looped chromatin organization at a developmentally regulated gene locus
.
Mol. Cell.
29
:
232
242
.
Kagey
,
M.H.
,
J.J.
Newman
,
S.
Bilodeau
,
Y.
Zhan
,
D.A.
Orlando
,
N.L.
van Berkum
,
C.C.
Ebmeier
,
J.
Goossens
,
P.B.
Rahl
,
S.S.
Levine
, et al
2010
.
Mediator and cohesin connect gene expression and chromatin architecture
.
Nature.
467
:
430
435
.
Kaneko
,
S.
,
J.
Son
,
R.
Bonasio
,
S.S.
Shen
, and
D.
Reinberg
.
2014
.
Nascent RNA interaction keeps PRC2 activity poised and in check
.
Genes Dev.
28
:
1983
1988
.
Kim
,
Y.H.
,
S.A.
Marhon
,
Y.
Zhang
,
D.J.
Steger
,
K.-J.
Won
, and
M.A.
Lazar
.
2018
.
Rev-erbα dynamically modulates chromatin looping to control circadian gene transcription
.
Science.
359
:
1274
1277
.
Kind
,
J.
,
L.
Pagie
,
H.
Ortabozkoyun
,
S.
Boyle
,
S.S.
de Vries
,
H.
Janssen
,
M.
Amendola
,
L.D.
Nolen
,
W.A.
Bickmore
, and
B.
van Steensel
.
2013
.
Single-cell dynamics of genome-nuclear lamina interactions
.
Cell.
153
:
178
192
.
Knight
,
J.D.
,
R.
Li
, and
M.
Botchan
.
1991
.
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
.
Krijger
,
P.H.L.
, and
W.
de Laat
.
2016
.
Regulation of disease-associated gene expression in the 3D genome
.
Nat. Rev. Mol. Cell Biol.
17
:
771
782
.
Kumaran
,
R.I.
, and
D.L.
Spector
.
2008
.
A genetic locus targeted to the nuclear periphery in living cells maintains its transcriptional competence
.
J. Cell Biol.
180
:
51
65
.
Kurukuti
,
S.
,
V.K.
Tiwari
,
G.
Tavoosidana
,
E.
Pugacheva
,
A.
Murrell
,
Z.
Zhao
,
V.
Lobanenkov
,
W.
Reik
, and
R.
Ohlsson
.
2006
.
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
.
Lai
,
F.
,
U.A.
Orom
,
M.
Cesaroni
,
M.
Beringer
,
D.J.
Taatjes
,
G.A.
Blobel
, and
R.
Shiekhattar
.
2013
.
Activating RNAs associate with Mediator to enhance chromatin architecture and transcription
.
Nature.
494
:
497
501
.
Lettice
,
L.A.
,
S.J.H.
Heaney
,
L.A.
Purdie
,
L.
Li
,
P.
de Beer
,
B.A.
Oostra
,
D.
Goode
,
G.
Elgar
,
R.E.
Hill
, and
E.
de Graaff
.
2003
.
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
.
Lewis
,
E.B.
1978
.
A gene complex controlling segmentation in Drosophila
.
Nature.
276
:
565
570
.
Li
,
H.-B.
,
M.
Müller
,
I.A.
Bahechar
,
O.
Kyrchanova
,
K.
Ohno
,
P.
Georgiev
, and
V.
Pirrotta
.
2011
.
Insulators, not Polycomb response elements, are required for long-range interactions between Polycomb targets in Drosophila melanogaster
.
Mol. Cell. Biol.
31
:
616
625
.
Li
,
H.-B.
,
K.
Ohno
,
H.
Gui
, and
V.
Pirrotta
.
2013
.
Insulators target active genes to transcription factories and polycomb-repressed genes to polycomb bodies
.
PLoS Genet.
9
:
e1003436
.
Lieberman-Aiden
,
E.
,
N.L.
van Berkum
,
L.
Williams
,
M.
Imakaev
,
T.
Ragoczy
,
A.
Telling
,
I.
Amit
,
B.R.
Lajoie
,
P.J.
Sabo
,
M.O.
Dorschner
, et al
2009
.
Comprehensive mapping of long-range interactions reveals folding principles of the human genome
.
Science.
326
:
289
293
.
Liu
,
Z.
,
D.R.
Scannell
,
M.B.
Eisen
, and
R.
Tjian
.
2011
.
Control of embryonic stem cell lineage commitment by core promoter factor, TAF3
.
Cell.
146
:
720
731
.
Lower
,
K.M.
,
J.R.
Hughes
,
M.
De Gobbi
,
S.
Henderson
,
V.
Viprakasit
,
C.
Fisher
,
A.
Goriely
,
H.
Ayyub
,
J.
Sloane-Stanley
,
D.
Vernimmen
, et al
2009
.
Adventitious changes in long-range gene expression caused by polymorphic structural variation and promoter competition
.
Proc. Natl. Acad. Sci. USA.
106
:
21771
21776
.
Lucas
,
J.S.
,
Y.
Zhang
,
O.K.
Dudko
, and
C.
Murre
.
2014
.
3D trajectories adopted by coding and regulatory DNA elements: first-passage times for genomic interactions
.
Cell.
158
:
339
352
.
Lupiáñez
,
D.G.
,
K.
Kraft
,
V.
Heinrich
,
P.
Krawitz
,
F.
Brancati
,
E.
Klopocki
,
D.
Horn
,
H.
Kayserili
,
J.M.
Opitz
,
R.
Laxova
, et al
2015
.
Disruptions of topological chromatin domains cause pathogenic rewiring of gene-enhancer interactions
.
Cell.
161
:
1012
1025
.
Mao
,
Y.S.
,
B.
Zhang
, and
D.L.
Spector
.
2011
.
Biogenesis and function of nuclear bodies
.
Trends Genet.
27
:
295
306
.
Meaburn
,
K.J.
, and
T.
Misteli
.
2007
.
Cell biology: chromosome territories
.
Nature.
445
:
379
781
.
Merika
,
M.
, and
D.
Thanos
.
2001
.
Enhanceosomes
.
Curr. Opin. Genet. Dev.
11
:
205
208
.
Mermet
,
J.
,
J.
Yeung
,
C.
Hurni
,
D.
Mauvoisin
,
K.
Gustafson
,
C.
Jouffe
,
D.
Nicolas
,
Y.
Emmenegger
,
C.
Gobet
,
P.
Franken
, et al
2018
.
Clock-dependent chromatin topology modulates circadian transcription and behavior
.
Genes Dev.
32
:
347
358
.
Michaelis
,
C.
,
R.
Ciosk
, and
K.
Nasmyth
.
1997
.
Cohesins: chromosomal proteins that prevent premature separation of sister chromatids
.
Cell.
91
:
35
45
.
Mifsud
,
B.
,
F.
Tavares-Cadete
,
A.N.
Young
,
R.
Sugar
,
S.
Schoenfelder
,
L.
Ferreira
,
S.W.
Wingett
,
S.
Andrews
,
W.
Grey
,
P.A.
Ewels
, et al
2015
.
Mapping long-range promoter contacts in human cells with high-resolution capture Hi-C
.
Nat. Genet.
47
:
598
606
.
Mitchell
,
J.A.
, and
P.
Fraser
.
2008
.
Transcription factories are nuclear subcompartments that remain in the absence of transcription
.
Genes Dev.
22
:
20
25
.
Moore
,
J.M.
,
N.A.
Rabaia
,
L.E.
Smith
,
S.
Fagerlie
,
K.
Gurley
,
D.
Loukinov
,
C.M.
Disteche
,
S.J.
Collins
,
C.J.
Kemp
,
V.V.
Lobanenkov
, and
G.N.
Filippova
.
2012
.
Loss of maternal CTCF is associated with peri-implantation lethality of Ctcf null embryos
.
PLoS One.
7
:
e34915
.
Moreau
,
P.
,
R.
Hen
,
B.
Wasylyk
,
R.
Everett
,
M.P.
Gaub
, and
P.
Chambon
.
1981
.
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
.
Mumbach
,
M.R.
,
A.J.
Rubin
,
R.A.
Flynn
,
C.
Dai
,
P.A.
Khavari
,
W.J.
Greenleaf
, and
H.Y.
Chang
.
2016
.
HiChIP: efficient and sensitive analysis of protein-directed genome architecture
.
Nat. Methods.
13
:
919
922
.
Mumbach
,
M.R.
,
A.T.
Satpathy
,
E.A.
Boyle
,
C.
Dai
,
B.G.
Gowen
,
S.W.
Cho
,
M.L.
Nguyen
,
A.J.
Rubin
,
J.M.
Granja
,
K.R.
Kazane
, et al
2017
.
Enhancer connectome in primary human cells identifies target genes of disease-associated DNA elements
.
Nat. Genet.
49
:
1602
1612
.
Murrell
,
A.
,
S.
Heeson
,
W.N.
Cooper
,
E.
Douglas
,
S.
Apostolidou
,
G.E.
Moore
,
E.R.
Maher
, and
W.
Reik
.
2004
a
.
An association between variants in the IGF2 gene and Beckwith-Wiedemann syndrome: interaction between genotype and epigenotype
.
Hum. Mol. Genet.
13
:
247
255
.
Murrell
,
A.
,
S.
Heeson
, and
W.
Reik
.
2004
b
.
Interaction between differentially methylated regions partitions the imprinted genes Igf2 and H19 into parent-specific chromatin loops
.
Nat. Genet.
36
:
889
893
.
Narendra
,
V.
,
P.P.
Rocha
,
D.
An
,
R.
Raviram
,
J.A.
Skok
,
E.O.
Mazzoni
, and
D.
Reinberg
.
2015
.
CTCF establishes discrete functional chromatin domains at the Hox clusters during differentiation
.
Science.
347
:
1017
1021
.
Nasmyth
,
K.
2001
.
Disseminating the genome: joining, resolving, and separating sister chromatids during mitosis and meiosis
.
Annu. Rev. Genet.
35
:
673
745
.
Németh
,
A.
,
A.
Conesa
,
J.
Santoyo-Lopez
,
I.
Medina
,
D.
Montaner
,
B.
Péterfia
,
I.
Solovei
,
T.
Cremer
,
J.
Dopazo
, and
G.
Längst
.
2010
.
Initial genomics of the human nucleolus
.
PLoS Genet.
6
:
e1000889
.
Nir
,
G.
,
I.
Farabella
,
C.P.
Estrada
,
C.G.
Ebeling
,
B.J.
Beliveau
,
H.M.
Sasaki
,
S.H.
Lee
,
S.C.
Nguyen
,
R.B.
McCole
,
S.
Chattoraj
, et al
2018
.
Walking along chromosomes with super-resolution imaging, contact maps, and integrative modeling
.
bioRxiv.
(Preprint posted July 28, 2018)
Nolis
,
I.K.
,
D.J.
McKay
,
E.
Mantouvalou
,
S.
Lomvardas
,
M.
Merika
, and
D.
Thanos
.
2009
.
Transcription factors mediate long-range enhancer-promoter interactions
.
Proc. Natl. Acad. Sci. USA.
106
:
20222
20227
.
Noordermeer
,
D.
,
M.R.
Branco
,
E.
Splinter
,
P.
Klous
,
W.
van Ijcken
,
S.
Swagemakers
,
M.
Koutsourakis
,
P.
van der Spek
,
A.
Pombo
, and
W.
de Laat
.
2008
.
Transcription and chromatin organization of a housekeeping gene cluster containing an integrated β-globin locus control region
.
PLoS Genet.
4
:
e1000016
.
Nora
,
E.P.
,
B.R.
Lajoie
,
E.G.
Schulz
,
L.
Giorgetti
,
I.
Okamoto
,
N.
Servant
,
T.
Piolot
,
N.L.
van Berkum
,
J.
Meisig
,
J.
Sedat
, et al
2012
.
Spatial partitioning of the regulatory landscape of the X-inactivation centre
.
Nature.
485
:
381
385
.
Nora
,
E.P.
,
A.
Goloborodko
,
A.-L.
Valton
,
J.H.
Gibcus
,
A.
Uebersohn
,
N.
Abdennur
,
J.
Dekker
,
L.A.
Mirny
, and
B.G.
Bruneau
.
2017
.
Targeted Degradation of CTCF Decouples Local Insulation of Chromosome Domains from Genomic Compartmentalization
.
Cell.
169
:
930
944.e22
.
Palstra
,
R.-J.
,
B.
Tolhuis
,
E.
Splinter
,
R.
Nijmeijer
,
F.
Grosveld
, and
W.
de Laat
.
2003
.
The beta-globin nuclear compartment in development and erythroid differentiation
.
Nat. Genet.
35
:
190
194
.
Palstra
,
R.-J.
,
M.
Simonis
,
P.
Klous
,
E.
Brasset
,
B.
Eijkelkamp
, and
W.
de Laat
.
2008
.
Maintenance of long-range DNA interactions after inhibition of ongoing RNA polymerase II transcription
.
PLoS One.
3
:
e1661
.
Parelho
,
V.
,
S.
Hadjur
,
M.
Spivakov
,
M.
Leleu
,
S.
Sauer
,
H.C.
Gregson
,
A.
Jarmuz
,
C.
Canzonetta
,
Z.
Webster
,
T.
Nesterova
, et al
2008
.
Cohesins functionally associate with CTCF on mammalian chromosome arms
.
Cell.
132
:
422
433
.
Patrinos
,
G.P.
,
M.
de Krom
,
E.
de Boer
,
A.
Langeveld
,
A.M.A.
Imam
,
J.
Strouboulis
,
W.
de Laat
, and
F.G.
Grosveld
.
2004
.
Multiple interactions between regulatory regions are required to stabilize an active chromatin hub
.
Genes Dev.
18
:
1495
1509
.
Peric-Hupkes
,
D.
,
W.
Meuleman
,
L.
Pagie
,
S.W.M.
Bruggeman
,
I.
Solovei
,
W.
Brugman
,
S.
Gräf
,
P.
Flicek
,
R.M.
Kerkhoven
,
M.
van Lohuizen
, et al
2010
.
Molecular maps of the reorganization of genome-nuclear lamina interactions during differentiation
.
Mol. Cell.
38
:
603
613
.
Phillips
,
J.E.
, and
V.G.
Corces
.
2009
.
CTCF: master weaver of the genome
.
Cell.
137
:
1194
1211
.
Phillips-Cremins
,
J.E.
,
M.E.G.
Sauria
,
A.
Sanyal
,
T.I.
Gerasimova
,
B.R.
Lajoie
,
J.S.K.
Bell
,
C.-T.
Ong
,
T.A.
Hookway
,
C.
Guo
,
Y.
Sun
, et al
2013
.
Architectural protein subclasses shape 3D organization of genomes during lineage commitment
.
Cell.
153
:
1281
1295
.
Picard
,
D.
, and
W.
Schaffner
.
1983
.
Correct transcription of a cloned mouse immunoglobulin gene in vivo
.
Proc. Natl. Acad. Sci. USA.
80
:
417
421
.
Pirrotta
,
V.
, and
H.-B.
Li
.
2012
.
A view of nuclear Polycomb bodies
.
Curr. Opin. Genet. Dev.
22
:
101
109
.
Pott
,
S.
, and
J.D.
Lieb
.
2015
.
What are super-enhancers?
Nat. Genet.
47
:
8
12
.
Quinodoz
,
S.A.
,
N.
Ollikainen
,
B.
Tabak
,
A.
Palla
,
J.M.
Schmidt
,
E.
Detmar
,
M.M.
Lai
,
A.A.
Shishkin
,
P.
Bhat
,
Y.
Takei
, et al
2018
.
Higher-Order Inter-chromosomal Hubs Shape 3D Genome Organization in the Nucleus
.
Cell.
174
:
744
757.e24
.
Ragoczy
,
T.
,
A.
Telling
,
T.
Sawado
,
M.
Groudine
, and
S.T.
Kosak
.
2003
.
A genetic analysis of chromosome territory looping: diverse roles for distal regulatory elements
.
Chromosome Res.
11
:
513
525
.
Ragoczy
,
T.
,
M.A.
Bender
,
A.
Telling
,
R.
Byron
, and
M.
Groudine
.
2006
.
The locus control region is required for association of the murine beta-globin locus with engaged transcription factories during erythroid maturation
.
Genes Dev.
20
:
1447
1457
.
Raj
,
A.
, and
A.
van Oudenaarden
.
2008
.
Nature, nurture, or chance: stochastic gene expression and its consequences
.
Cell.
135
:
216
226
.
Rao
,
S.S.P.
,
M.H.
Huntley
,
N.C.
Durand
,
E.K.
Stamenova
,
I.D.
Bochkov
,
J.T.
Robinson
,
A.L.
Sanborn
,
I.
Machol
,
A.D.
Omer
,
E.S.
Lander
, and
E.L.
Aiden
.
2014
.
A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping
.
Cell.
159
:
1665
1680
.
Rao
,
S.S.P.
,
S.-C.
Huang
,
B.
Glenn St Hilaire
,
J.M.
Engreitz
,
E.M.
Perez
,
K.-R.
Kieffer-Kwon
,
A.L.
Sanborn
,
S.E.
Johnstone
,
G.D.
Bascom
,
I.D.
Bochkov
, et al
2017
.
Cohesin Loss Eliminates All Loop Domains
.
Cell.
171
:
305
320.e24
.
Recillas-Targa
,
F.
,
M.J.
Pikaart
,
B.
Burgess-Beusse
,
A.C.
Bell
,
M.D.
Litt
,
A.G.
West
,
M.
Gaszner
, and
G.
Felsenfeld
.
2002
.
Position-effect protection and enhancer blocking by the chicken beta-globin insulator are separable activities
.
Proc. Natl. Acad. Sci. USA.
99
:
6883
6888
.
Reddy
,
K.L.
,
J.M.
Zullo
,
E.
Bertolino
, and
H.
Singh
.
2008
.
Transcriptional repression mediated by repositioning of genes to the nuclear lamina
.
Nature.
452
:
243
247
.
Respuela
,
P.
,
M.
Nikolić
,
M.
Tan
,
P.
Frommolt
,
Y.
Zhao
,
J.
Wysocka
, and
A.
Rada-Iglesias
.
2016
.
Foxd3 Promotes Exit from Naive Pluripotency through Enhancer Decommissioning and Inhibits Germline Specification
.
Cell Stem Cell.
18
:
118
133
.
Rollins
,
R.A.
,
P.
Morcillo
, and
D.
Dorsett
.
1999
.
Nipped-B, a Drosophila homologue of chromosomal adherins, participates in activation by remote enhancers in the cut and Ultrabithorax genes
.
Genetics.
152
:
577
593
.
Rowley
,
M.J.
,
M.H.
Nichols
,
X.
Lyu
,
M.
Ando-Kuri
,
I.S.M.
Rivera
,
K.
Hermetz
,
P.
Wang
,
Y.
Ruan
, and
V.G.
Corces
.
2017
.
Evolutionarily Conserved Principles Predict 3D Chromatin Organization
.
Mol. Cell.
67
:
837
852.e7
.
Rubio
,
E.D.
,
D.J.
Reiss
,
P.L.
Welcsh
,
C.M.
Disteche
,
G.N.
Filippova
,
N.S.
Baliga
,
R.
Aebersold
,
J.A.
Ranish
, and
A.
Krumm
.
2008
.
CTCF physically links cohesin to chromatin
.
Proc. Natl. Acad. Sci. USA.
105
:
8309
8314
.
Sadoni
,
N.
,
B.-S.
Targosz
,
A.
Englmann
,
S.
Fesser
,
J.
Koch
,
D.
Schindelhauer
, and
D.
Zink
.
2008
.
Transcription-dependent spatial arrangements of CFTR and conserved adjacent loci are not conserved in human and murine nuclei
.
Chromosoma.
117
:
381
397
.
Sagai
,
T.
,
H.
Masuya
,
M.
Tamura
,
K.
Shimizu
,
Y.
Yada
,
S.
Wakana
,
Y.
Gondo
,
T.
Noda
, and
T.
Shiroishi
.
2004
.
Phylogenetic conservation of a limb-specific, cis-acting regulator of Sonic hedgehog (Shh)
.
Mamm. Genome.
15
:
23
34
.
Sanborn
,
A.L.
,
S.S.P.
Rao
,
S.-C.
Huang
,
N.C.
Durand
,
M.H.
Huntley
,
A.I.
Jewett
,
I.D.
Bochkov
,
D.
Chinnappan
,
A.
Cutkosky
,
J.
Li
, et al
2015
.
Chromatin extrusion explains key features of loop and domain formation in wild-type and engineered genomes
.
Proc. Natl. Acad. Sci. USA.
112
:
E6456
E6465
.
Sanyal
,
A.
,
B.R.
Lajoie
,
G.
Jain
, and
J.
Dekker
.
2012
.
The long-range interaction landscape of gene promoters
.
Nature.
489
:
109
113
.
Schaaf
,
C.A.
,
Z.
Misulovin
,
M.
Gause
,
A.
Koenig
,
D.W.
Gohara
,
A.
Watson
, and
D.
Dorsett
.
2013
.
Cohesin and polycomb proteins functionally interact to control transcription at silenced and active genes
.
PLoS Genet.
9
:
e1003560
.
Schmidt
,
D.
,
P.C.
Schwalie
,
M.D.
Wilson
,
B.
Ballester
,
A.
Gonçalves
,
C.
Kutter
,
G.D.
Brown
,
A.
Marshall
,
P.
Flicek
, and
D.T.
Odom
.
2012
.
Waves of retrotransposon expansion remodel genome organization and CTCF binding in multiple mammalian lineages
.
Cell.
148
:
335
348
.
Schnappauf
,
O.
,
S.
Beyes
,
A.
Dertmann
,
V.
Freihen
,
P.
Frey
,
S.
Jägle
,
K.
Rose
,
T.
Michoel
,
R.
Grosschedl
, and
A.
Hecht
.
2016
.
Enhancer decommissioning by Snail1-induced competitive displacement of TCF7L2 and down-regulation of transcriptional activators results in EPHB2 silencing
.
Biochim. Biophys. Acta.
1859
:
1353
1367
.
Schoenfelder
,
S.
,
M.
Furlan-Magaril
,
B.
Mifsud
,
F.
Tavares-Cadete
,
R.
Sugar
,
B.-M.
Javierre
,
T.
Nagano
,
Y.
Katsman
,
M.
Sakthidevi
,
S.W.
Wingett
, et al
2015
.
The pluripotent regulatory circuitry connecting promoters to their long-range interacting elements
.
Genome Res.
25
:
582
597
.
Schwartz
,
Y.B.
, and
G.
Cavalli
.
2017
.
Three-Dimensional Genome Organization and Function in Drosophila
.
Genetics.
205
:
5
24
.
Schwartz
,
Y.B.
, and
V.
Pirrotta
.
2007
.
Polycomb silencing mechanisms and the management of genomic programmes
.
Nat. Rev. Genet.
8
:
9
22
.
Schwarzer
,
W.
,
N.
Abdennur
,
A.
Goloborodko
,
A.
Pekowska
,
G.
Fudenberg
,
Y.
Loe-Mie
,
N.A.
Fonseca
,
W.
Huber
,
C.
H Haering
,
L.
Mirny
, and
F.
Spitz
.
2017
.
Two independent modes of chromatin organization revealed by cohesin removal
.
Nature.
551
:
51
56
.
Sexton
,
T.
,
E.
Yaffe
,
E.
Kenigsberg
,
F.
Bantignies
,
B.
Leblanc
,
M.
Hoichman
,
H.
Parrinello
,
A.
Tanay
, and
G.
Cavalli
.
2012
.
Three-dimensional folding and functional organization principles of the Drosophila genome
.
Cell.
148
:
458
472
.
Sigrist
,
C.J.
, and
V.
Pirrotta
.
1997
.
Chromatin insulator elements block the silencing of a target gene by the Drosophila polycomb response element (PRE) but allow trans interactions between PREs on different chromosomes
.
Genetics.
147
:
209
221
.
Simon
,
J.
,
A.
Chiang
,
W.
Bender
,
M.J.
Shimell
, and
M.
O’Connor
.
1993
.
Elements of the Drosophila bithorax complex that mediate repression by Polycomb group products
.
Dev. Biol.
158
:
131
144
.
Song
,
S.-H.
,
C.
Hou
, and
A.
Dean
.
2007
.
A positive role for NLI/Ldb1 in long-range beta-globin locus control region function
.
Mol. Cell.
28
:
810
822
.
Soshnikova
,
N.
,
T.
Montavon
,
M.
Leleu
,
N.
Galjart
, and
D.
Duboule
.
2010
.
Functional analysis of CTCF during mammalian limb development
.
Dev. Cell.
19
:
819
830
.
Splinter
,
E.
,
H.
Heath
,
J.
Kooren
,
R.-J.
Palstra
,
P.
Klous
,
F.
Grosveld
,
N.
Galjart
, and
W.
de Laat
.
2006
.
CTCF mediates long-range chromatin looping and local histone modification in the β-globin locus
.
Genes Dev.
20
:
2349
2354
.
Struhl
,
G.
1981
.
A gene product required for correct initiation of segmental determination in Drosophila
.
Nature.
293
:
36
41
.
Su
,
W.
,
S.
Jackson
,
R.
Tjian
, and
H.
Echols
.
1991
.
DNA looping between sites for transcriptional activation: self-association of DNA-bound Sp1
.
Genes Dev.
5
:
820
826
.
Sundaram
,
V.
,
Y.
Cheng
,
Z.
Ma
,
D.
Li
,
X.
Xing
,
P.
Edge
,
M.P.
Snyder
, and
T.
Wang
.
2014
.
Widespread contribution of transposable elements to the innovation of gene regulatory networks
.
Genome Res.
24
:
1963
1976
.
Symmons
,
O.
,
V.V.
Uslu
,
T.
Tsujimura
,
S.
Ruf
,
S.
Nassari
,
W.
Schwarzer
,
L.
Ettwiller
, and
F.
Spitz
.
2014
.
Functional and topological characteristics of mammalian regulatory domains
.
Genome Res.
24
:
390
400
.
Symmons
,
O.
,
L.
Pan
,
S.
Remeseiro
,
T.
Aktas
,
F.
Klein
,
W.
Huber
, and
F.
Spitz
.
2016
.
The Shh Topological Domain Facilitates the Action of Remote Enhancers by Reducing the Effects of Genomic Distances
.
Dev. Cell.
39
:
529
543
.
Szabo
,
Q.
,
D.
Jost
,
J.-M.
Chang
,
D.I.
Cattoni
,
G.L.
Papadopoulos
,
B.
Bonev
,
T.
Sexton
,
J.
Gurgo
,
C.
Jacquier
,
M.
Nollmann
, et al
2018
.
TADs are 3D structural units of higher-order chromosome organization in Drosophila
.
Sci. Adv.
4
:
eaar8082
.
Tang
,
Z.
,
O.J.
Luo
,
X.
Li
,
M.
Zheng
,
J.J.
Zhu
,
P.
Szalaj
,
P.
Trzaskoma
,
A.
Magalska
,
J.
Wlodarczyk
,
B.
Ruszczycki
, et al
2015
.
CTCF-Mediated Human 3D Genome Architecture Reveals Chromatin Topology for Transcription
.
Cell.
163
:
1611
1627
.
Therizols
,
P.
,
R.S.
Illingworth
,
C.
Courilleau
,
S.
Boyle
,
A.J.
Wood
, and
W.A.
Bickmore
.
2014
.
Chromatin decondensation is sufficient to alter nuclear organization in embryonic stem cells
.
Science.
346
:
1238
1242
.
Tolhuis
,
B.
,
R.-J.
Palstra
,
E.
Splinter
,
F.
Grosveld
, and
W.
de Laat
.
2002
.
Looping and interaction between hypersensitive sites in the active beta-globin locus
.
Mol. Cell.
10
:
1453
1465
.
Vakoc
,
C.R.
,
D.L.
Letting
,
N.
Gheldof
,
T.
Sawado
,
M.A.
Bender
,
M.
Groudine
,
M.J.
Weiss
,
J.
Dekker
, and
G.A.
Blobel
.
2005
.
Proximity among distant regulatory elements at the beta-globin locus requires GATA-1 and FOG-1
.
Mol. Cell.
17
:
453
462
.
van Koningsbruggen
,
S.
,
M.
Gierlinski
,
P.
Schofield
,
D.
Martin
,
G.J.
Barton
,
Y.
Ariyurek
,
J.T.
den Dunnen
, and
A.I.
Lamond
.
2010
.
High-resolution whole-genome sequencing reveals that specific chromatin domains from most human chromosomes associate with nucleoli
.
Mol. Biol. Cell.
21
:
3735
3748
.
van Steensel
,
B.
, and
A.S.
Belmont
.
2017
.
Lamina-Associated Domains: Links with Chromosome Architecture, Heterochromatin, and Gene Repression
.
Cell.
169
:
780
791
.
Vernimmen
,
D.
,
M.
De Gobbi
,
J.A.
Sloane-Stanley
,
W.G.
Wood
, and
D.R.
Higgs
.
2007
.
Long-range chromosomal interactions regulate the timing of the transition between poised and active gene expression
.
EMBO J.
26
:
2041
2051
.
Vietri Rudan
,
M.
,
C.
Barrington
,
S.
Henderson
,
C.
Ernst
,
D.T.
Odom
,
A.
Tanay
, and
S.
Hadjur
.
2015
.
Comparative Hi-C reveals that CTCF underlies evolution of chromosomal domain architecture
.
Cell Reports.
10
:
1297
1309
.
Wang
,
L.
,
J.L.
Brown
,
R.
Cao
,
Y.
Zhang
,
J.A.
Kassis
, and
R.S.
Jones
.
2004
.
Hierarchical recruitment of polycomb group silencing complexes
.
Mol. Cell.
14
:
637
646
.
Wang
,
S.
,
J.-H.
Su
,
B.J.
Beliveau
,
B.
Bintu
,
J.R.
Moffitt
,
C.-T.
Wu
, and
X.
Zhuang
.
2016
.
Spatial organization of chromatin domains and compartments in single chromosomes
.
Science.
353
:
598
602
.
Watrin
,
E.
,
F.J.
Kaiser
, and
K.S.
Wendt
.
2016
.
Gene regulation and chromatin organization: relevance of cohesin mutations to human disease
.
Curr. Opin. Genet. Dev.
37
:
59
66
.
Watson
,
L.A.
,
X.
Wang
,
A.
Elbert
,
K.D.
Kernohan
,
N.
Galjart
, and
N.G.
Bérubé
.
2014
.
Dual effect of CTCF loss on neuroprogenitor differentiation and survival
.
J. Neurosci.
34
:
2860
2870
.
Weintraub
,
A.S.
,
C.H.
Li
,
A.V.
Zamudio
,
A.A.
Sigova
,
N.M.
Hannett
,
D.S.
Day
,
B.J.
Abraham
,
M.A.
Cohen
,
B.
Nabet
,
D.L.
Buckley
, et al
2017
.
YY1 Is a Structural Regulator of Enhancer-Promoter Loops
.
Cell.
171
:
1573
1588.e28
.
Wendt
,
K.S.
,
K.
Yoshida
,
T.
Itoh
,
M.
Bando
,
B.
Koch
,
E.
Schirghuber
,
S.
Tsutsumi
,
G.
Nagae
,
K.
Ishihara
,
T.
Mishiro
, et al
2008
.
Cohesin mediates transcriptional insulation by CCCTC-binding factor
.
Nature.
451
:
796
801
.
West
,
A.G.
,
M.
Gaszner
, and
G.
Felsenfeld
.
2002
.
Insulators: many functions, many mechanisms
.
Genes Dev.
16
:
271
288
.
West
,
A.G.
,
S.
Huang
,
M.
Gaszner
,
M.D.
Litt
, and
G.
Felsenfeld
.
2004
.
Recruitment of histone modifications by USF proteins at a vertebrate barrier element
.
Mol. Cell.
16
:
453
463
.
Whyte
,
W.A.
,
S.
Bilodeau
,
D.A.
Orlando
,
H.A.
Hoke
,
G.M.
Frampton
,
C.T.
Foster
,
S.M.
Cowley
, and
R.A.
Young
.
2012
.
Enhancer decommissioning by LSD1 during embryonic stem cell differentiation
.
Nature.
482
:
221
225
.
Wijchers
,
P.J.
,
G.
Geeven
,
M.
Eyres
,
A.J.
Bergsma
,
M.
Janssen
,
M.
Verstegen
,
Y.
Zhu
,
Y.
Schell
,
C.
Vermeulen
,
E.
de Wit
, and
W.
de Laat
.
2015
.
Characterization and dynamics of pericentromere-associated domains in mice
.
Genome Res.
25
:
958
969
.
Wijchers
,
P.J.
,
P.H.L.
Krijger
,
G.
Geeven
,
Y.
Zhu
,
A.
Denker
,
M.J.A.M.
Verstegen
,
C.
Valdes-Quezada
,
C.
Vermeulen
,
M.
Janssen
,
H.
Teunissen
, et al
2016
.
Cause and Consequence of Tethering a SubTAD to Different Nuclear Compartments
.
Mol. Cell.
61
:
461
473
.
Wijgerde
,
M.
,
F.
Grosveld
, and
P.
Fraser
.
1995
.
Transcription complex stability and chromatin dynamics in vivo
.
Nature.
377
:
209
213
.
Williams
,
A.
,
C.G.
Spilianakis
, and
R.A.
Flavell
.
2010
.
Interchromosomal association and gene regulation in trans
.
Trends Genet.
26
:
188
197
.
Williamson
,
I.
,
L.A.
Lettice
,
R.E.
Hill
, and
W.A.
Bickmore
.
2016
.
Shh and ZRS enhancer colocalisation is specific to the zone of polarising activity
.
Development.
143
:
2994
3001
.
Würtele
,
H.
, and
P.
Chartrand
.
2006
.
Genome-wide scanning of HoxB1-associated loci in mouse ES cells using an open-ended Chromosome Conformation Capture methodology
.
Chromosome Res.
14
:
477
495
.
Yadon
,
A.N.
,
B.N.
Singh
,
M.
Hampsey
, and
T.
Tsukiyama
.
2013
.
DNA looping facilitates targeting of a chromatin remodeling enzyme
.
Mol. Cell.
50
:
93
103
.
Yoon
,
Y.S.
,
S.
Jeong
,
Q.
Rong
,
K.-Y.
Park
,
J.H.
Chung
, and
K.
Pfeifer
.
2007
.
Analysis of the H19ICR insulator
.
Mol. Cell. Biol.
27
:
3499
3510
.
Zhang
,
Y.
,
C.-H.
Wong
,
R.Y.
Birnbaum
,
G.
Li
,
R.
Favaro
,
C.Y.
Ngan
,
J.
Lim
,
E.
Tai
,
H.M.
Poh
,
E.
Wong
, et al
2013
.
Chromatin connectivity maps reveal dynamic promoter-enhancer long-range associations
.
Nature.
504
:
306
310
.
Zhao
,
H.
, and
A.
Dean
.
2004
.
An insulator blocks spreading of histone acetylation and interferes with RNA polymerase II transfer between an enhancer and gene
.
Nucleic Acids Res.
32
:
4903
4919
.
This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see http://www.rupress.org/terms/). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 4.0 International license, as described at https://creativecommons.org/licenses/by-nc-sa/4.0/).