A tour of 3D genome with a focus on CTCF

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A tour of 3D genome with a focus on CTCF Diane C. Wang 1 , William Wang 1 , Linlin Zhang 1 , Xiangdong Wang ∗ Zhongshan Hospital Institute of Clinical Science, Zhongshan Hospital, Fudan University Medical School, Shanghai Institute of Clinical Bioinformatics Shanghai, China

a r t i c l e

i n f o

Article history: Received 12 July 2018 Accepted 17 July 2018 Available online xxx Keywords: 3D genome CTCF DNA binding TADs Degradation Wapl

a b s t r a c t The complex three-dimensional (3D) structure of the genome plays critical roles in the maintenance of genome stability, organization, and dynamics and in regulation of gene expression for understanding molecular mechanisms and diseases. Chromatin maintains biological functions and transcriptional activities through long distance interaction and interactions between loops and enhancers-promoters. We firstly overview the architecture and biology of chromatin and loops, topologically associated domains (TADs) and interactions, and compartments and functions. We specifically focus on CCCTC-binding factor (CTCF) in 3D genome organization and function to furthermore understand the significance of CTCF biology, transcriptional regulations, interactions with cohesin, roles in DNA binding, influences of CTCF degradation, and communication with wings-apart like (Wapl) protein. We also summarize the advanced single cell approaches to further monitor dynamics of CTCF functions and structures in the maintenance of 3D genome organization and function at single cell level. © 2018 Published by Elsevier Ltd.

Contents 1. 2. 3. 4. 5.

6. 7. 8.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Interactions with chromatin and loops . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Functions of topologically associated domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Biology and characteristics of compartments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Roles of CTCF in 3D genome organization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.1. CTCF Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.2. Transcriptional regulations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.3. Interactions with cohesin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.4. Mechanisms of CTCF binding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 5.5. Influences of CTCF degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Communication with Wapl . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Single cell approaches in CTCF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00 Conclusion and perspectives. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .00 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 00

1. Introduction The complex three-dimensional (3D) structure of the genome plays critical roles in the maintenance of genome stability, organization, and dynamics and in regulation of gene expression for understanding molecular mechanisms and diseases [1–4]. Chro-

∗ Corresponding author. E-mail address: [email protected] (X. Wang). 1 Authors contribute to this article equally as the first author.

mosomes are separated into chromosome territories in the nucleus whereas the chromosome itself is partitioned into compartments followed by topologically associated domains (TADs), loops, and chromatin itself [5] (Fig. 1). This process is achieved partially with the help of the high concentration of proteins, such as CTCF and cohesion, in the nucleus [6]. The present article takes a rapid tour through the architecture and biology of 3D genome and related important elements, e.g. chromatin and loops, TADs and interactions, and compartments and functions. We specifically focus on CCCTC-binding factor (CTCF), one of the most important regulators in 3D genome organization and function, by addressing CTCF biology, transcriptional regulations, interactions with cohesin, roles

https://doi.org/10.1016/j.semcdb.2018.07.020 1084-9521/© 2018 Published by Elsevier Ltd.

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Fig. 1. Chromosome territories. On a chromosomal level, it is separated into chromosome territories whereas the chromosome itself is partitioned into compartments A/B, followed by topologically associated domains (TAD), DNA loops and finally the chromatin itself. Approximately 30% of the loops bring together promoters and enhancers.

in DNA binding, influences of CTCF degradation, and communication with wings-apart like (Wapl) protein. We also summarize the advanced single cell approaches to further monitor dynamics of CTCF functions and structures in the maintenance of 3D genome organization and function at single cell level.

tioning as a cellular memory [14]. Genes are more selected in a cluster than just preferentially colocalized, although further studies are required to clarify the process. Faunucchi et al. found that removing even a single gene from the cluster changed the transcription of co-associated genes, which would argue for a higher degree of co-ordination between the genes [15].

2. Interactions with chromatin and loops Chromatin refers to DNA wrapped around histones and other DNA-binding proteins to form 10 nm thick chromatin fibers [1]. Chromatin has long distance interactions due to pairs of loci that have stronger interaction than with any loci in between them, and can be within and across topological domains [7]. Approximately 30% of loops bring together promoters and enhancers [8], which is related with transcriptional activity and gene expression [9]. About 66% of active promoters contact the nearest enhancer, 30% bypass at least one, and 4% contact the closest one in one direction even though the opposite direction holds a closer one. Of those, 90% interact with at least one more distant enhancer, form complex and multi-interaction networks (Fig. 2). This is in contradiction from the previous theory that enhancers only interact with the closest promoters [10]. Javierre et al. found 17,500 interactions between promoters and promoter interacting regions, with a median of 4 interactions per promoter. Of those, about 50% interacted with one promoter and 10% with four or more, indicating that multiple regulatory elements contribute to gene expression. From an evolutionary point of view, this could be a safety feature to mitigate the potential pathological effect if one enhancer is affected [11], although Sagai et al. demonstrated that the limb-specific sonic hedgehog enhancers control the gene expression and limb degeneration [12]. Inter-gene promoter loop interactions result from communications between transcription factors (TFs) and the associated genes, to form gene clusters as specialized nuclear hot spots for efficient transcription [13]. The mechanisms by which TF binding pattern is inherited following DNA synthesis and condensation of chromosomes remain unclear. All clusters have cohesin, while loss of cohesin decreases DNA accessibility and TF binding to clusters. Cohesin enables the re-establishment of TF clusters through func-

3. Functions of topologically associated domains TADs are partitions of chromosomes with a high frequency of interactions between loci. Loci located in different TADs have low interaction frequency even if they happen to situated next to each other [16]. This indicates a regulatory effect in constraining the region that an enhancer can affect. The insertion of a regulatory sensor into random genomic positions demonstrated distinct selfinteracting blocks in the genome [17] and had specific repeats and CTCF-binding sites with tissue-specific expression potentials [18,19]. Although exact number and size of the domains remains difficult to pinpoint, numbers vary from 2200 domains with a median size of 880 kb in mice [16] to anything between 4000 to 9000 domains with sizes between 40 kb to 3 Mb in humans [8]. TADs are divided into smaller sub-TADs of a few hundred kb [20] or “contact domains” between 10–100 kb long [8]. TADs and sub-TADs have interaction loops with varied length and strength identified on Hi-C maps [5]. Certain characteristics of domains are enriched with CTCG, active transcription marks, nascent transcripts, repeat elements, housekeeping genes, and transfer RNA [8,16]. TADs play a functional role in the human genome and may be a distinguished hierarchy level as TADs scales differ from other [21,49]. Functional features such as CTCF clustering and transcriptional coregulation during differentiation are correlated with TAD scales [22]. In a clinical context, disrupted domains were linked with distinct limb malformations and cancer secondary to pathogenic gene regulation [23,24]. The deletion of TADs in mitotic chromosomes indicates that the genomic structure appears to be fluid and changes during the cell cycle [23]. This would account for the high cell-to-cell variability demonstrated in single-cell Hi-C and fluorescence in situ hybridization (FISH) studies [25,26].

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Fig. 2. Active promoters and enhancer. 66% of active promoters contact the nearest enhancer, but that 30% will bypass at least one and the remaining 4% contact the closest one in one direction even though the opposite direction holds a closer one. Moreover, regardless of which of the above category they belonged to, 90% will interact with at least one more distant enhancer, thus showing the formation of complex and multi-way interaction networks.

4. Biology and characteristics of compartments The concept of partitioning each chromosome into two compartments was introduced after noticing a plaid-like pattern of large alternating block of enriched and depleted interaction frequencies [27]. The compartments are referred to as compartment A, more open, accessible, and actively transcribed, and B, a relatively compact state of chromatin. The loci are associated with higher expression and chromatin accessibility in compartment A, instead of higher tendency for close spatial localization as in compartment B. Open and closed chromatin domains reside in different spatial compartments [28]. Those two compartments are further divided into A1-2 and B1-4 on basis of chromatin interaction patterns [29]. Sub-compartments A1 and A2 share highly expressed genes and chromatin marks characteristic of active chromatin, such as H3K79me2, H3K27ac, and H3K36me3. A2 has longer genes, lower GC content, and completes replication later than A1. Compartment B1, B2, B3, and B4, are located in the periphery of the nucleus or to nucleoli. B1 has more epigenetic mark indicative of facultative heterochromatin, B2 contains mainly pericentromeric heterochromatin, B3 is enriched at the nuclear lamina but depleted at the nucleolus-associated domains, and B4 is only present on chromosome 19 and spans 11Mb [8,28]. The fractal globule, as a proposed polymer model for the genome compartment architecture, includes interphase DNA to self-organize into a long-lived and non-equilibrium conformation [30,31]. An untangled polymer forms series of small globules, which yields a “beads on a string” appearance, and acts as monomers in further rounds of crumpling [32]. The contact probability obtained from Hi-C data is close to that obtained from analysis of the fractal globule [27], similar to scaling of 3D distance between predicted and data reported by 3D FISH [33]. 5. Roles of CTCF in 3D genome organization 5.1. CTCF Biology The CTCF protein, an architectural protein, is regarded as the “master weaver” of the genome [6]. The protein is made up of 11 zin. fingers (ZF), stabilized by zinc ions binding to cysteine and histidine residues, which reside in a central DNA-binding domain, and

is flanked by unstructured N- and C-terminal domains [34]. The “CTCF code” hypothesis speculates that CTCF recognizes diverse DNA sequences through combinatorial usage of its 11 ZFs [35]. CTCF regulates gene expression through enhancer blocking activity by vertebral insulators, blocks long-range interactions, and separates active from silent chromatins [6,36,37]. CTCF contributes to the link between genome spatial organization and function through interactions between regulatory sequences [33]. The presence of 11 ZF domain-compromising residues 266–579 and the 3 ZF domain-compromising residues 402–494 changes DNA architecture from relaxed morphology in the control samples to compact circular complexes, meshes, and networks [38]. However, some contradictory results are about CTCF importance in the genome organization process. For example, CTCF depletion mildly affected TADs [29], probably due to incomplete loos of CTCF [33]. Complete loss of CTCF causes loss of insulation between neighboring TADs, while chromatin loops remained, although with reduced strength. However, separating function of active CTCF from silent chromatin was not observed [17,37]. Using auxin-inducible degron techniques to deplete CTCF in mouse embryonic stem cells, LADassociated domains were found disintegrable, but other domains remained stable [39]. 5.2. Transcriptional regulations Transcriptional regulation in three-dimensional space is an important part of mechanisms by which methyl-sensitive CTCF binding acts as functional enhancer blocking. The imprinted control region (ICR) is methylated on the paternal allele, and eliminates CTCF binding and ICR-mediated insulation, leading to functional communication between promoters and enhancers and up-regulation of insulin-like growth-factor 2 (Igf2) expressions [36]. ICR is unmethylated on maternal alleles and CTCF is bound, while Igf2 promoters are prevented from accessing the enhancer downstream of H19 [39]. Because of the promoter methylation, H19 expression is suppressed on the parent allele and disconnected from the CTCF binding of the methylated ICR. This explains the potential direct role of CTCF in transcriptional regulation, that is, the promotion of proximal placement may protect genes from methylation-dependent silencing [36]. De novo methylation of CpG islands in the promoter region is an epigenetic feature of gene

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silencing in cancer [37]. DNA methylation reduces the expression of prostaglandin-endoperoxide synthase 2 (PTGS2), which is an inducible, rate-limiting enzyme. Abnormal metabolites produced by mutant isocitrate dehydrogenase 1 (IDH1) may drive the production of glioma mainly by changing 3D conformation of DNA [40]. Reduction of CTCF binding at a domain boundary, equivalent with loss of insulation between topological domains and abnormal gene activation, allows a constitutive enhancer to interact aberrantly with PDGFRA. In IDH1 mutant cells the CpG dinucleotide in the CTCF presents higher methylation, while demethylation could restore CTCF insulator function of mediated insulation [40] (Fig. 4). 5.3. Interactions with cohesin Cohesin is another architectural protein complex and consists of a large ring-shaped molecule to encircle two strands of DNA, to form the complex with CTCF/and target hypomethylated DNA. Cohesin was initially recognized as a component for achieving cohesion between two sister chromosomes [12,41]. Cohesin coupled together with CTCF play critical roles in transcription [42,43] and works independently from CTCF, when loaded to promoter and enhancer elements or in estrogen-regulated transcription [44,45]. The chromatin looping mediated by CTCF/cohesin down-regulates prostaglandin synthesis. In the vicinity of CpG islands associated with PTGS2, the number of CTCF/cohesin complexes is abundant. PTGS locus forms chromatin loops through the methylation-sensitive binding of the complex, of which links are eliminated by DNA methylation (Fig. 3). 5.4. Mechanisms of CTCF binding The exact mechanism by which CTCF binds to many degenerate DNA sequences remains unclear. Study on the structure and mechanics of CTCF and DNA binding demonstrated that ZF3-7 have high specificity in the affinity to a core sequence derived from human chromosome 5 through forming base-specific contacts [46]. Out of 15-bp core motifs, base pairs in the 2nd, 3rd, 6th, 7th, 8th, 10th, 11th, and 12th positions showed the highest likelihood of having conserved bps with H-bonds between bases of the top strand and the ZF residue. The remaining and variable bps shared weak Hbonds, water-mediated H-bonds, hydrophobic interaction, or gap between the protein and DNA, with the core sequence. The ability of ZFs to adapt to various sequences resulted from the hydrogen bonds with variable bases to replace a G:C base pair instead of an A:T base pair at the 3rd position, without altering the binding affinity [46]. Methylation of CTCF loop anchor sites is speculated to disable CTCF binding, interaction with enhancers, and expression of inappropriate genes outside of the loops [47,48]. Process of targeting catalytically inactive Cas9 to two CTCF binding sites induced de novo methylation [49]. There is a clear correlation between decreased bound CTCF to targeted genome sites and increased interaction frequency between super-enhancers in the miR290 with activated neighboring gene, NLR family pyrin domain containing 12. This was a result of a diminished insulator effect of CTCF and increased interaction frequency between nearby, previously insulated, super-enhancers, and genes [48]. Cytosine methylations in mammalian cells occur at positions 2 and 12, followed by guanine or adenine to form CpG or CpA [50]. 40% of variable CTCF binding is linked to differential methylation and occur mostly at those two sites [51]. Changes of a methyl group were at the C5 atom of cytosine in position 2 and drastically decreased the affinity for the oligo due to blocking of ZF7 D451. An increased C2 methylation decreases CTCF binding, while increasing C12 methylation improves CTCF binding. CTCF recognizes sequences with high variability in the 15bp core sequence motif through ZF residues and forms multiple

conformations through establishing versatile H-bonds with some bases [46]. 5.5. Influences of CTCF degradation Auxin-inducible degron (AID) system targets CTCF stop codons and introduces an amino acid variant of AID tags using enhanced green fluorescent protein cassettes to reduce CTCF levels [52–54]. Proteasome-dependent degradation affects the bond formation between trypanosome infection response 1 F-box protein and AID. Sub-cloning of CTCF reduced cells in AID systems can be prevented by auxin-related degradation. Similar observations were made in complete CTCF knockout stem cells and additionally, cell proliferation decreased after prolonged CTCF depletion [52–55]. CTCF is crucial for interactions of the X-inactivation centre (Xic), evidenced by the finding that the anchorage of CTCF interactions and 3C carbon copy 5C peaks were non-existent in CTCP depleted cells [56]. Disruption of folding and resting astrocytes results in misfolding at the Xic locus. Nora et al. recently published a milestone study to understand the consequences of CTCF degradation and molecular mechanisms by which the degradation contributes to CTCF function [52]. TAD boundaries could be responsible for protection of promoters, which interact with neighboring enhancers, and mis regulated gene promoters are frequently found in close proximity to loop anchors. Inter-TAD 3D distance, equivalent to intra-TAD distances, is reduced by CTCF loss and compacting sequences in different TADs are responsible for insulation reduction. Furthermore, chromatin compaction is not affected by the presence of CTCF, although absence of CTCF makes linear genomic coordinates more preferable for measurement of 3D distances. The 3D nucleus space has increased TAD boundaries consistent with reduced insulation. There is little known about the control of chromatin basal packaging and the segregation of A and B compartments. There are no alterations of genomic location of transitions between the A- and B- compartments and the scaling of contact frequencies, and minimal reductions of compartmentalization strength, in the absence of CTCF [52]. Lack of CTCF directly impacts population of TADs due to failure of chromatin communication and a dosedependent curve. Severe or complete reduction of CTCF needs to occur before organizational chromosomal defects occur. CTCF may be responsible for transcription promotion by manipulating nucleosomes to limit occlusion promotion and regulation of responsive genes and by affecting tracking processes not related to chromatin loop accumulation [52]. CTCF mediates enhancer-blocked insulation interactions at a genome scale by influencing boundaries of TAD. CTCF absence causes up-regulation of certain genes which previously was isolated from neighbor enhancers [47,57]. In reversible models of CTCF degradation, a subset of genes sustains irreversible changes of transcriptional signals post auxin wash off. It is possible that CTCF affects and contributes to regulate some factors, which are involved in positive feedback mechanism. 6. Communication with Wapl The Wapl protein plays an important role in the dynamic interaction of cohesin with chromatin by controlling the interlocking gate mechanism at structural maintenance of chromosomes 1 and 3, as well as SCC1 [58–60]. Absence of Wapl reduces cohesin turnover on chromatin, increases DNA and TAD interaction, and increases median loop lengths [61,62], suggesting that cohesin turnover controls the intra-TAD interactions and loop lengths. Wapl activation maintains the balance of loop formation, while a deficit of Wapl results in tandem loop formation in upregulated sites. An increased frequency of peaks in the 5 and 3 intersection boundary is responsible for enlarging smaller chromatin loops into larger

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Fig. 3. CTCF in chromatin organization of mouse H19/Igf2 locus. The maternally expressed noncoding H19 gene is located approximately 90 kb downstream from the gene encoding Igf2 that is expressed exclusively from the paternal allele. The ICR about 2 kb upstream of H19 contains four CTCF-binding sites and is essential for regulation of the entire locus. Differentially methylated regions (DMRs), such as DMR1 upstream of Igf2 promoters) and DMR2 within Igf2 exon 6. ICR is methylated on the paternal allele, which eliminates CTCF binding and ICR-mediated insulation, leading to functional communication between promoters and enhancers, thereby activating Igf2 expressions. ICR is unmethylated on maternal alleles and CTCF is bound, while Igf2 promoters are prevented from accessing the enhancer downstream of H19. 2b. CTCF / Cohesin stabilizes the long term chromatin interaction by interacting with other CTCF / cohesin sites. But DNA methylation of the PTGS2 CpG island blocks the links among CTCF/cohesin-binding sites at the PTGS2 locus by restraining the enrichment of CTCF/cohesin at these regions.

Fig. 4. In IDH wild-type cells, intact CTCF act as insulating role of oncogene, while disruption of the boundary by IDH mutation and hypermethylation allows a potent constitutive enhancer to interact abnormally with PDGFRA and upregulate it. IDH mutation lead to CTCF CpG methylation and removing the CTCF binding, which activate the oncogene PDGFRA. The interaction between FIP1L1 enhancer and PDGFRA promoter in IDH mutant cells is stronger than that between FIP1L1 enhancer and FIP1L1 promoter, and result PDGFRA highly expressed. Methyltransferase inhibitor 5-azacytidine treated could reduce methylation of the CTCF motif, increased CTCF binding and downregulated PDGFRA expression.

loops and TADs are a polyclonal collection of loops in the middle of separate CTCF sites [61]. CTCF associated and non-associated cohesin sites increases frequency of structural maintenance of chromosomes 1 binding signals in WAPL deficient cells. Nuclear compartmentalization is regulated by the amount of cohesin present on chromatin to enable chromosome flexibility and the removal of Wapl/SSC4 alters the topology of the entire chromosome [61]. Insertion of gene traps in the genes encoding subunit of SCC2 and SCC4 cohesion loader complex subunits did not impair

the proliferation of Wapl deficient cells suggesting that increased cohesin and DNA interaction time offsets the absence of Wapl. SCC2 C-terminal is integral for formation of Vermicelli chromosome and in SCC2 deficient cells SCC4 is deleted on cohesins [61].The finding that Wapl and SCC4 deficient cells had similar stability in DNAbound cohesin stability shows no requirement for SCC4 presence during cohesin loading. Deletion of CTCF or Wapl alone has significantly more canonical cohesin-bonding sites as compared to deletion of both CTCF and Wapl, which has larger cohesin-bound

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areas in actively transcribed genes. Wapl can be translocated with cohesin and the absence of sororin could initiate cohesin release [63,64]. 7. Single cell approaches in CTCF Single cell sequencing, systems biology, and intelligent models provide a new vision to understand heterogeneity, origin, phenome, drug sensitivity, and transcriptional regulation at single cell level [65–68]. The integration of gene sequencing, transcriptional regulation, and molecular transomics with phenomes can present deeper and broader figures of each gene and molecule [69,70]. Potential mechanisms by which single-cell diversity occurs are associated with the binding of CTCF with the cohesin complex subunit Rad21 to highly conserved DNA sequences through alterations of transcriptionally active Pcdh␣ promoters, isoform expression, spaced pair of CTCF/Rad21 binding sites, and distant enhancer elements [71]. Stevens et al. made an outstanding investigation on 3D structures of entire genomes, genome folding, and each TADs and loops at a scale < 100 kb in single cell level using data from the 3C method [72]. CTCF/cohesin loops interact with at least one end of the loop in or very near to compartment A, of which functional heterogeneity exist among cells. For example, CTCF/cohesin loops are formed in about 1/3 cells and contact boundaries vary 12–62% of the cells [72]. This particular study showed strong evidence that TADs and CTCF/cohesin loops form in partial cells and have clear changes of structure dynamics and variations. Ren et al. integrated single-cell flow cytometry and single-molecule RNA-FISH assays with gene editing technology and initially found that CTCF could stabilize interaction of enhancer and promoter to maintain minimal variations of gene expression [73]. CTCF binding sites have a close interlinking with enhancers within TADs and are activated with the up-regulation of corresponding enhancers. This particular study provides the important evidence that CTCF controls target gene expression through regulation of long-range promoter-enhancer interaction. 8. Conclusion and perspectives The complex three-dimensional (3D) structure of genome plays critical roles in maintenance of genome stability, organization, and dynamics and in regulation of gene expression for understanding molecular mechanisms and diseases. Chromatin maintains biological functions and transcriptional activities through long distance interaction and interactions between loops and enhancers-promoters. We firstly overview architecture and biology of chromatin and loops, TADs and interactions, and compartments and functions. We specially focus on CTCF in 3D genome organization and function to furthermore understand significance of CTCF biology, transcriptional regulations, interactions with cohesin, roles in DNA binding, influences of CTCF degradation, and communication with Wapl protein. We also summarize the advanced single cell approaches to further monitor dynamics of CTCF functions and structures in maintenance of 3D genome organization and function at single cell level. The progress in our knowledge of the 3D genome has been growing exponentially and the multitude of loop types, shapes, and functions within the chromatin is more complex than we expect. The exact mechanism by which loops fold and shaped as well as the interplay between genes and TFs remains to be furthermore explored. There are urgent needs identify and validate CTCF-associated biomarkers and targets with specificity of biological function and transcriptional regulation, as well as disease phenome, phase, duration, severity, and response to therapy [74–80]. Systems heterogeneity is an emerging science to design

new strategies for drug therapy, understand roles of epigenetics and target molecules as well as immune microenvironment of target cells, and illustrate dynamics and patterns of heterogeneityassociated metabolisms [81–87]. With development of single cell technology, heterogeneity of dynamic biology and regulation of CTCF and associated elements can be deeply investigated. The complexity of studying the human genome lies in the vastness of components that all require further discovery and clarification.

References [1] P. Szalaj, D. Plewczynski, Three-dimensional organization and dynamics of the genome, Cell Biol. Toxicol. (2018), http://dx.doi.org/10.1007/s10565-0189428-y, ahead of print. [2] R. Li, Y. Liu, Y. Hou, J. Gan, P. Wu, C. Li, 3D genome and its disorganization in diseases, Cell Biol. Toxicol. (2018), http://dx.doi.org/10.1007/s10565-0189430-4, ahead of print. [3] T. Ma, L. Chen, M. Shi, J. Niu, X. Zhang, X. Yang, K. Zhanghao, M. Wang, P. Xi, D. Jin, M. Zhang, J. Gao, Developing novel methods to image and visualize 3D genomes, Cell Biol. Toxicol. (2018), http://dx.doi.org/10.1007/s10565-0189427-z, ahead of print. [4] T. Terabayashi, K. Hanada, Genome instability syndromes caused by impaired DNA repair and aberrant DNA damage responses, Cell Biol. Toxicol. (2018), http://dx.doi.org/10.1007/s10565-018-9429-x. ˜ S. Mundlos, Structural variation in the 3D [5] M. Spielmann, D.G. Lupiánez, genome, Nat. Rev. Genet. 19 (2018) 453–467, http://dx.doi.org/10.1038/ s41576-018-0007-0. [6] J.E. Phillips, V.G. Corces, CTCF: master weaver of the genome, Cell 137 (2009) 1194–1211, http://dx.doi.org/10.1016/j.cell.2009.06.001. [7] S. Kadauke, G.A. Blobel, Chromatin loops in gene regulation, Biochim. Biophys. Acta - Gene Regul. Mech. 1789 (2009) 17–25, http://dx.doi.org/10.1016/j. bbagrm.2008.07.002. [8] S.S.P. Rao, 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, E.L. Aiden, A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping, Cell 159 (2014) 1665–1680, http://dx.doi.org/10.1016/j.cell.2014.11. 021. [9] B. Bonev, N. Mendelson Cohen, Q. Szabo, L. Fritsch, G.L. Papadopoulos, Y. Lubling, X. Xu, X. Lv, J.-P. Hugnot, A. Tanay, G. Cavalli, 3D. Multiscale, Genome rewiring during mouse neural development, Cell 171 (2017) 557–572, http:// dx.doi.org/10.1016/j.cell.2017.09.043, e24. [10] S. Schoenfelder, M. Furlan-Magaril, B. Mifsud, F. Tavares-Cadete, R. Sugar, B.-M. Javierre, T. Nagano, Y. Katsman, M. Sakthidevi, S.W. Wingett, E. Dimitrova, A. Dimond, L.B. Edelman, S. Elderkin, K. Tabbada, E. Darbo, S. Andrews, B. Herman, A. Higgs, E. LeProust, C.S. Osborne, J.A. Mitchell, N.M. Luscombe, P. Fraser, The pluripotent regulatory circuitry connecting promoters to their long-range interacting elements, Genome Res. 25 (2015) 582–597, http://dx.doi.org/10.1101/gr.185272.114. [11] B.M. Javierre, O.S. Burren, S.P. Wilder, R. Kreuzhuber, S.M. Hill, S. Sewitz, J. Cairns, S.W. Wingett, C. Várnai, M.J. Thiecke, F. Burden, S. Farrow, A.J. Cutler, K. Rehnström, K. Downes, L. Grassi, M. Kostadima, P. Freire-Pritchett, F. Wang, H.G. BLUEPRINT Consortium, H.G. Stunnenberg, J.A. Todd, D.R. Zerbino, O. Stegle, W.H. Ouwehand, M. Frontini, C. Wallace, M. Spivakov, P. Fraser, B. Kim, N. Sharifi, E.M. Janssen-Megens, M.-L. Yaspo, M. Linser, A. Kovacsovics, L. Clarke, D. Richardson, A. Datta, P. Flicek, Lineage-specific genome architecture links enhancers and non-coding disease variants to target gene promoters, Cell 167 (2016) 1369–1384, http://dx.doi.org/10.1016/j.cell.2016.09.037, e19. [12] T. Sagai, M. Hosoya, Y. Mizushina, M. Tamura, T. Shiroishi, Elimination of a long-range cis-regulatory module causes complete loss of limb-specific Shh expression and truncation of the mouse limb, Development 132 (2005) 797–803, http://dx.doi.org/10.1242/dev.01613. [13] S. Schoenfelder, T. Sexton, L. Chakalova, N.F. Cope, A. Horton, S. Andrews, S. Kurukuti, J.A. Mitchell, D. Umlauf, D.S. Dimitrova, C.H. Eskiw, Y. Luo, C.-L. Wei, Y. Ruan, J.J. Bieker, P. Fraser, Preferential associations between co-regulated genes reveal a transcriptional interactome in erythroid cells, Nat. Genet. 42 (2010) 53–61, http://dx.doi.org/10.1038/ng.496. [14] J. Yan, M. Enge, T. Whitington, K. Dave, J. Liu, I. Sur, B. Schmierer, A. Jolma, T. Kivioja, M. Taipale, J. Taipale, Transcription factor binding in human cells occurs in dense clusters formed around cohesin anchor sites, Cell 154 (2013) 801–813, http://dx.doi.org/10.1016/j.cell.2013.07.034. [15] S. Fanucchi, Y. Shibayama, S. Burd, M.S. Weinberg, M.M. Mhlanga, Chromosomal contact permits transcription between coregulated genes, Cell 155 (2013) 606–620, http://dx.doi.org/10.1016/j.cell.2013.09.051. [16] J.R. Dixon, S. Selvaraj, F. Yue, A. Kim, Y. Li, Y. Shen, M. Hu, J.S. Liu, B. Ren, Topological domains in mammalian genomes identified by analysis of chromatin interactions, Nature 485 (2012) 376–380, http://dx.doi.org/10. 1038/nature11082. [17] J. Zuin, J.R. Dixon, M.I.J.A. van der Reijden, Z. Ye, P. Kolovos, R.W.W. Brouwer, M.P.C. van de Corput, H.J.G. van de Werken, T.A. Knoch, W.F.J. van IJcken, F.G. Grosveld, B. Ren, K.S. Wendt, Cohesin and CTCF differentially affect chromatin architecture and gene expression in human cells, Proc. Natl. Acad. Sci. U. S. A. (2014), http://dx.doi.org/10.1073/pnas.1317788111.

Please cite this article in press as: D.C. Wang, et al., A tour of 3D genome with a focus on CTCF, Semin Cell Dev Biol (2018), https://doi.org/10.1016/j.semcdb.2018.07.020

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ARTICLE IN PRESS D.C. Wang et al. / Seminars in Cell & Developmental Biology xxx (2018) xxx–xxx

[18] S. Ruf, O. Symmons, V.V. Uslu, D. Dolle, C. Hot, L. Ettwiller, F. Spitz, Large-scale analysis of the regulatory architecture of the mouse genome with a transposon-associated sensor, Nat. Genet. 43 (2011) 379–386, http://dx.doi. org/10.1038/ng.790. [19] O. Symmons, V.V. Uslu, T. Tsujimura, S. Ruf, S. Nassari, W. Schwarzer, L. Ettwiller, F. Spitz, Functional and topological characteristics of mammalian regulatory domains, Genome Res. 24 (2014) 390–400, http://dx.doi.org/10. 1101/gr.163519.113. [20] J.E. Phillips-Cremins, 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, M.J. Bland, W. Wagstaff, S. Dalton, T.C. McDevitt, R. Sen, J. Dekker, J. Taylor, V.G. Corces, Architectural protein subclasses shape 3D organization of genomes during lineage commitment, Cell 153 (2013) 1281–1295, http://dx.doi.org/10.1016/j.cell.2013.04.053. [21] Z. Jowhar, P.R. Gudla, S. Shachar, D. Wangsa, J.L. Russ, G. Pegoraro, T. Ried, A. Raznahan, T. Misteli, HiCTMap: detection and analysis of chromosome territory structure and position by high-throughput imaging, Methods 142 (2018) 30–38, http://dx.doi.org/10.1016/j.ymeth.2018.01.013. [22] Y. Zhan, L. Mariani, I. Barozzi, E.G. Schulz, N. Blüthgen, M. Stadler, G. Tiana, L. Giorgetti, Reciprocal insulation analysis of Hi-C data shows that TADs represent a functionally but not structurally privileged scale in the hierarchical folding of chromosomes, Genome Res. 27 (2017) 479–490, http:// dx.doi.org/10.1101/gr.212803.116. ˜ K. Kraft, V. Heinrich, P. Krawitz, F. Brancati, E. Klopocki, D. Horn, [23] D.G. Lupiánez, H. Kayserili, J.M. Opitz, R. Laxova, F. Santos-Simarro, B. Gilbert-Dussardier, L. Wittler, M. Borschiwer, S.A. Haas, M. Osterwalder, M. Franke, B. Timmermann, J. Hecht, M. Spielmann, A. Visel, S. Mundlos, Disruptions of topological chromatin domains cause pathogenic rewiring of gene-enhancer interactions, Cell 161 (2015) 1012–1025, http://dx.doi.org/10.1016/j.cell.2015.04.004. [24] D. Hnisz, 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, J. Reddy, D. Borges-Rivera, T.I. Lee, R. Jaenisch, M.H. Porteus, J. Dekker, R.A. Young, Activation of proto-oncogenes by disruption of chromosome neighborhoods, Science 351 (2016) 1454–1458, http://dx.doi.org/10.1126/science.aad9024. [25] N. Naumova, M. Imakaev, G. Fudenberg, Y. Zhan, B.R. Lajoie, L.A. Mirny, J. Dekker, Organization of the mitotic chromosome, Science (80-.) 342 (2013) 948–953, http://dx.doi.org/10.1126/science.1236083. [26] T. Nagano, Y. Lubling, C. Várnai, C. Dudley, W. Leung, Y. Baran, N. Mendelson Cohen, S. Wingett, P. Fraser, A. Tanay, Cell-cycle dynamics of chromosomal organization at single-cell resolution, Nature 547 (2017) 61–67, http://dx.doi. org/10.1038/nature23001. [27] L. Giorgetti, R. Galupa, E.P. Nora, T. Piolot, F. Lam, J. Dekker, G. Tiana, E. Heard, Predictive polymer modeling reveals coupled fluctuations in chromosome conformation and transcription, Cell 157 (2014) 950–963, http://dx.doi.org/ 10.1016/j.cell.2014.03.025. [28] E. Lieberman-Aiden, N.L. van Berkum, L. Williams, M. Imakaev, T. Ragoczy, A. Telling, I. Amit, B.R. Lajoie, P.J. Sabo, M.O. Dorschner, R. Sandstrom, B. Bernstein, M.A. Bender, M. Groudine, A. Gnirke, J. Stamatoyannopoulos, L.A. Mirny, E.S. Lander, J. Dekker, Comprehensive mapping of long-range interactions reveals folding principles of the human genome, Science 326 (2009) 289–293, http://dx.doi.org/10.1126/science.1181369. [29] M.T. Mawhinney, R. Liu, F. Lu, J. Maksimoska, K. Damico, R. Marmorstein, P.M. Lieberman, B. Urbanc, CTCF-induced circular DNA complexes observed by atomic force microscopy, J. Mol. Biol. 430 (2018) 759–776, http://dx.doi.org/ 10.1016/j.jmb.2018.01.012. ´ V. Heinrich, D.M. Ibrahim, C. Paliou, M. [30] G. Andrey, R. Schöpflin, I. Jerkovic, Hochradel, B. Timmermann, S. Haas, M. Vingron, S. Mundlos, Characterization of hundreds of regulatory landscapes in developing limbs reveals two regimes of chromatin folding, Genome Res. 27 (2017) 223–233, http://dx.doi.org/10. 1101/gr.213066.116. [31] A.I. Grosberg, S.K. Nechaev, E.I. Shakhnovich, [The role of topological limitations in the kinetics of homopolymer collapse and self-assembly of biopolymers], Biofizika. 33 (n.d.) 247–53. http://www.ncbi.nlm.nih.gov/ pubmed/3390477 (Accessed July 10, 2018). [32] A.Y. Grosberg, Y. Rabin, S. Havlin, A. Neer, Self-similarity in the structure of DNA - what are introns, Biofizika 38 (1993) 75–83. [33] J. Mateos-Langerak, M. Bohn, W. de Leeuw, O. Giromus, E.M.M. Manders, P.J. Verschure, M.H.G. Indemans, H.J. Gierman, D.W. Heermann, R. van Driel, S. Goetze, Spatially confined folding of chromatin in the interphase nucleus, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 3812–3817, http://dx.doi.org/10.1073/ pnas.0809501106. [34] S.S. Roy, A.K. Mukherjee, S. Chowdhury, Insights about genome function from spatial organization of the genome, Hum. Genomics 12 (2018) 8, http://dx. doi.org/10.1186/s40246-018-0140-z. [35] S.R. Martinez, J.L. Miranda, CTCF terminal segments are unstructured, Protein Sci. 19 (2010) 1110–1116, http://dx.doi.org/10.1002/pro.367. [36] D. Schmidt, P.C. Schwalie, M.D. Wilson, B. Ballester, Â. Gonc¸alves, C. Kutter, G.D. Brown, A. Marshall, P. Flicek, D.T. Odom, Waves of retrotransposon expansion remodel genome organization and CTCF binding in multiple mammalian lineages, Cell 148 (2012) 335–348, http://dx.doi.org/10.1016/j. cell.2011.11.058. [37] C. Hou, H. Zhao, K. Tanimoto, A. Dean, CTCF-dependent enhancer-blocking by alternative chromatin loop formation, Proc. Natl. Acad. Sci. U. S. A. 105 (2008) 20398–20403, http://dx.doi.org/10.1073/pnas.0808506106. [38] V. Narendra, P.P. Rocha, D. An, R. Raviram, J.A. Skok, E.O. Mazzoni, D. Reinberg, CTCF establishes discrete functional chromatin domains at the Hox clusters

[39]

[40]

[41]

[42]

[43]

[44]

[45]

[46]

[47]

[48]

[49]

[50]

[51]

[52]

[53]

[54]

[55]

[56]

[57]

[58]

[59]

7

during differentiation, Science 347 (2015) 1017–1021, http://dx.doi.org/10. 1126/science.1262088. N. Kubo, H. Ishii, D. Gorkin, F. Meitinger, X. Xiong, R. Fang, T. Liu, Z. Ye, B. Li, J. Dixon, A. Desai, H. Zhao, B. Ren, Preservation of chromatin organization after acute loss of CTCF in mouse embryonic stem cells, BioRxiv (2017), http://dx. doi.org/10.1101/118737, ahead of print. R. Sherwood, T.S. Takahashi, P.V. Jallepalli, Sister acts: coordinating DNA replication and cohesion establishment, Genes Dev. (2010), http://dx.doi.org/ 10.1101/gad.1976710. G.G. Galli, M. Carrara, C. Francavilla, K. Honnens de Lichtenberg, J.V. Olsen, R.A. Calogero, A.H. Lund, Genomic and proteomic analyses of Prdm5 reveal interactions with insulator binding proteins in embryonic stem cells, Mol. Cell. Biol. 33 (2013) 4504–4516, http://dx.doi.org/10.1128/MCB.00545-13. K.S. Wendt, J.M. Peters, How cohesin and CTCF cooperate in regulating gene expression, Chromosom. Res. 17 (2009) 201–214, http://dx.doi.org/10.1007/ s10577-008-9017-7. M.H. Kagey, 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, D.J. Taatjes, J. Dekker, R.A. Young, Mediator and cohesin connect gene expression and chromatin architecture, Nature 467 (2010) 430–435, http://dx.doi.org/10.1038/ nature09380. D. Schmidt, P.C. Schwalie, C.S. Ross-Innes, A. Hurtado, G.D. Brown, J.S. Carroll, P. Flicek, D.T. Odom, A CTCF-independent role for cohesin in tissue-specific transcription, Genome Res. 20 (2010) 578–588, http://dx.doi.org/10.1101/gr. 100479.109. V. Ramani, X. Deng, R. Qiu, K.L. Gunderson, F.J. Steemers, C.M. Disteche, W.S. Noble, Z. Duan, J. Shendure, Massively multiplex single-cell Hi-C, Nat. Methods 14 (2017) 263–720, http://dx.doi.org/10.1038/nmeth.4155. H. Hashimoto, D. Wang, J.R. Horton, X. Zhang, V.G. Corces, X. Cheng, Structural basis for the versatile and methylation-dependent binding of CTCF to DNA, Mol. Cell 66 (2017) 711–720, http://dx.doi.org/10.1016/j.molcel.2017.05.004. J.M. Dowen, Z.P. Fan, D. Hnisz, G. Ren, B.J. Abraham, L.N. Zhang, A.S. Weintraub, J. Schuijers, T.I. Lee, K. Zhao, R.A. Young, Control of cell identity genes occurs in insulated neighborhoods in mammalian chromosomes, Cell 159 (2014) 374–387, http://dx.doi.org/10.1016/j.cell.2014.09.030. A.C. Bell, G. Felsenfeld, Methylation of a CTCF-dependent boundary controls imprinted expression of the Igf2 gene, Nature 405 (2000) 482–485, http://dx. doi.org/10.1038/35013100. X.S. Liu, H. Wu, X. Ji, Y. Stelzer, X. Wu, S. Czauderna, J. Shu, D. Dadon, R.A. Young, R. Jaenisch, Editing DNA methylation in the mammalian genome, Cell 167 (2016) 233–247, http://dx.doi.org/10.1016/j.cell.2016.08.056, e17. R. Lister, M. Pelizzola, R.H. Dowen, R.D. Hawkins, G. Hon, J. Tonti-Filippini, J.R. Nery, L. Lee, Z. Ye, Q.M. Ngo, L. Edsall, J. Antosiewicz-Bourget, R. Stewart, V. Ruotti, A.H. Millar, J.A. Thomson, B. Ren, J.R. Ecker, Human DNA methylomes at base resolution show widespread epigenomic differences, Nature 462 (2009) 315–322, http://dx.doi.org/10.1038/nature08514. H. Wang, M.T. Maurano, H. Qu, K.E. Varley, J. Gertz, F. Pauli, K. Lee, T. Canfield, M. Weaver, R. Sandstrom, R.E. Thurman, R. Kaul, R.M. Myers, J.A. Stamatoyannopoulos, Widespread dplasticity in CTCF occupancy linked to DNA methylation, Genome Res. 22 (2012) 1680–1688, http://dx.doi.org/10. 1101/gr.136101.111. E.P. Nora, A. Goloborodko, A.L. Valton, J.H. Gibcus, A. Uebersohn, N. Abdennur, J. Dekker, L.A. Mirny, B.G. Bruneau, Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization, Cell 169 (2017) 930–944, http://dx.doi.org/10.1016/j. cell.2017.05.004. K. Nishimura, T. Fukagawa, H. Takisawa, T. Kakimoto, M. Kanemaki, An auxin-based degron system for the rapid depletion of proteins in nonplant cells, Nat. Methods. 6 (2009) 917–922, http://dx.doi.org/10.1038/nmeth.1401. M. Morawska, H.D. Ulrich, An expanded tool kit for the auxin-inducible degron system in budding yeast, Yeast 30 (2013) 341–351, http://dx.doi.org/ 10.1002/yea.2967. F. Sleutels, W. Soochit, M. Bartkuhn, H. Heath, S. Dienstbach, P. Bergmaier, V. Franke, M. Rosa-Garrido, S. Van De Nobelen, L. Caesar, M. Van Der Reijden, J.C. Bryne, W. Van Ijcken, J.A. Grootegoed, M.D. Delgado, B. Lenhard, R. Renkawitz, F. Grosveld, N. Galjart, The male germ cell gene regulator CTCFL is functionally different from CTCF and binds CTCF-like consensus sites in a nucleosome composition-dependent manner, Epigenet. Chrom. 5 (2012) 8, http://dx.doi. org/10.1186/1756-8935-5-8. J. Dostie, T.A. Richmond, R.A. Arnaout, R.R. Selzer, W.L. Lee, T.A. Honan, E.D. Rubio, A. Krumm, J. Lamb, C. Nusbaum, R.D. Green, J. Dekker, Chromosome conformation capture carbon copy (5C): a massively parallel solution for mapping interactions between genomic elements, Genome Res. 16 (2006) 1299–1309, http://dx.doi.org/10.1101/gr.5571506. B. Doyle, G. Fudenberg, M. Imakaev, L.A. Mirny, Chromatin loops as allosteric modulators of enhancer-promoter interactions, PLoS Comput. Biol. 10 (2014) e1003867, http://dx.doi.org/10.1371/journal.pcbi.1003867. S. Kueng, B. Hegemann, B.H. Peters, J.J. Lipp, A. Schleiffer, K. Mechtler, J.M. Peters, Wapl controls the dynamic association of cohesin with chromatin, Cell 127 (2006) 955–967, http://dx.doi.org/10.1016/j.cell.2006.09.040. F. Beckouët, M. Srinivasan, M.B. Roig, K.L. Chan, J.C. Scheinost, P. Batty, B. Hu, N. Petela, T. Gligoris, A.C. Smith, L. Strmecki, B.D. Rowland, K. Nasmyth, Releasing activity disengages cohesin’s Smc3/Scc1 interface in a process blocked by acetylation, Mol. Cell 61 (2016) 563–574, http://dx.doi.org/10. 1016/j.molcel.2016.01.026.

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[60] Y. Murayama, F. Uhlmann, DNA entry into and exit out of the cohesin ring by an interlocking gate mechanism, Cell 163 (2015) 1628–1640, http://dx.doi. org/10.1016/j.cell.2015.11.030. ˜ [61] J.H.I. Haarhuis, R.H. van der Weide, V.A. Blomen, J.O. Yánez-Cuna, M. Amendola, M.S. van Ruiten, P.H.L. Krijger, H. Teunissen, R.H. Medema, B. van Steensel, T.R. Brummelkamp, E. de Wit, B.D. Rowland, The cohesin release factor WAPL restricts chromatin loop extension, Cell 169 (2017) 693–707, http://dx.doi.org/10.1016/j.cell.2017.04.013. [62] J.D.P. Rhodes, J.H.I. Haarhuis, J.B. Grimm, B.D. Rowland, L.D. Lavis, K.A. Nasmyth, Cohesin can remain associated with chromosomes during DNA replication, Cell Rep 20 (2017) 2749–2755, http://dx.doi.org/10.1016/j.celrep. 2017.08.092. [63] T. Nishiyama, R. Ladurner, J. Schmitz, E. Kreidl, A. Schleiffer, V. Bhaskara, M. Bando, K. Shirahige, A.A. Hyman, K. Mechtler, J.M. Peters, Sororin mediates sister chromatid cohesion by antagonizing Wapl, Cell 143 (2010) 737–749, http://dx.doi.org/10.1016/j.cell.2010.10.031. [64] G.A. Busslinger, R.R. Stocsits, P. Van Der Lelij, E. Axelsson, A. Tedeschi, N. Galjart, J.M. Peters, Cohesin is positioned in mammalian genomes by transcription, CTCF and Wapl, Nature 544 (2017) 503–507, http://dx.doi.org/ 10.1038/nature22063. [65] W. Wang, D. Gao, X. Wang, Can single-cell RNA sequencing crack the mystery of cells? Cell Biol. Toxicol. 34 (2018) 1–6, http://dx.doi.org/10.1007/s10565017-9404-y. [66] W. Wang, B. Zhu, X. Wang, Dynamic phenotypes: illustrating a single-cell odyssey, Cell Biol. Toxicol. 33 (2017) 423–427, http://dx.doi.org/10.1007/ s10565-017-9400-2. [67] W. Wang, X. Wang, Single-cell CRISPR screening in drug resistance, Cell Biol. Toxicol. 33 (2017) 207–210, http://dx.doi.org/10.1007/s10565-017-9396-7. [68] M.P. Chu, J. Kriangkum, C.P. Venner, I. Sandhu, J. Hewitt, A.R. Belch, L.M. Pilarski, Addressing heterogeneity of individual blood cancers: the need for single cell analysis, Cell Biol. Toxicol. 33 (2017) 83–97, http://dx.doi.org/10. 1007/s10565-016-9367-4. [69] Y. Zeng, X. Chen, H. Gao, X. Wang, An artificial intelligent single cell is part of the cell dream world, Cell Biol. Toxicol. 34 (2018) 247–249, http://dx.doi.org/ 10.1007/s10565-018-9433-1. [70] X. Wang, Clinical trans-omics: an integration of clinical phenomes with molecular multiomics, Cell Biol. Toxicol. 34 (2018) 163–166, http://dx.doi. org/10.1007/s10565-018-9431-3. [71] K. Monahan, N.D. Rudnick, P.D. Kehayova, F. Pauli, K.M. Newberry, R.M. Myers, T. Maniatis, Role of CCCTC binding factor (CTCF) and cohesin in the generation of single-cell diversity of Protocadherin- gene expression, Proc. Natl. Acad. Sci. 109 (2006) 9125–9130, http://dx.doi.org/10.1073/pnas.1205074109. [72] T.J. Stevens, D. Lando, S. Basu, L.P. Atkinson, Y. Cao, S.F. Lee, M. Leeb, K.J. Wohlfahrt, W. Boucher, A. O’Shaughnessy-Kirwan, J. Cramard, A.J. Faure, M. Ralser, E. Blanco, L. Morey, M. Sansó, M.G.S. Palayret, B. Lehner, L. Di Croce, A. Wutz, B. Hendrich, D. Klenerman, E.D. Laue, 3D structures of individual mammalian genomes studied by single-cell Hi-C, Nature 544 (2017) 59–64, http://dx.doi.org/10.1038/nature21429.

[73] G. Ren, W. Jin, K. Cui, J. Rodrigez, G. Hu, Z. Zhang, D.R. Larson, K. Zhao, CTCF-mediated enhancer-promoter interaction Is a critical regulator of cell-to-cell variation of Gene expression, Mol. Cell 67 (2017) 1049–1058, http://dx.doi.org/10.1016/j.molcel.2017.08.026. [74] L. Shi, B. Zhu, M. Xu, X. Wang, Selection of AECOPD-specific immunomodulatory biomarkers by integrating genomics and proteomics with clinical informatics, Cell Biol. Toxicol. 34 (April (2)) (2018) 109–123, http://dx.doi.org/10.1007/s10565-017-9405-x. [75] Y. Kawamura, J. Takouda, K. Yoshimoto, K. Nakashima, New aspects of glioblastoma multiforme revealed by similarities between neural and glioblastoma stem cells, Cell Biol. Toxicol. 31 (January) (2018), http://dx.doi. org/10.1007/s10565-017-9420-y. [76] D. Long, T. Yu, X. Chen, Y.1 Liao, X. Lin, RNAi targeting STMN alleviates the resistance to taxol and collectively contributes to down regulate the malignancy of NSCLC cells in vitro and in vivo, Cell Biol. Toxicol. 34 (February (1)) (2018) 7–21, http://dx.doi.org/10.1007/s10565-017-9398-5. [77] D. Wu, X. Wang, H. Sun, The role of mitochondria in cellular toxicity as a potential drug target, Cell Biol. Toxicol. 34 (April (2)) (2018) 87–91, http://dx. doi.org/10.1007/s10565-018-9425-1. [78] X. Liu, J. Wu, History, applications, and challenges of immune repertoire research, Cell Biol. Toxicol. 27 (February) (2018), http://dx.doi.org/10.1007/ s10565-018-9426-0. [79] L. Shi, N. Dong, D. Ji, X. Huang, Z. Ying, X. Wang, C. Chen, Lipopolysaccharide-induced CCN1 production enhances interleukin-6 secretion in bronchial epithelial cells, Cell Biol. Toxicol. 34 (February (1)) (2018) 39–49, http://dx.doi.org/10.1007/s10565-017-9401-1. [80] M. Xu, X. Wang, Critical roles of mucin-1 in sensitivity of lung cancer cells to tumor necrosis factor-alpha and dexamethasone, Cell Biol. Toxicol. 33 (August (4)) (2017) 361–371, http://dx.doi.org/10.1007/s10565-017-9393-x. [81] D.C. Wang, W. Wang, B. Zhu, X. Wang, Lung cancer heterogeneity and New strategies for drug therapy, Annu. Rev. Pharmacol. Toxicol. 6 (January (58)) (2018) 531–546, http://dx.doi.org/10.1146/annurev-pharmtox-010716104523. [82] D.C. Wang, X. Wang, Systems heterogeneity: an integrative way to understand cancer heterogeneity, Semin. Cell Dev. Biol. 64 (April) (2017) 1–4. [83] N. Dong, L. Shi, D.C. Wang, C. Chen, X. Wang, Role of epigenetics in lung cancer heterogeneity and clinical implication, Semin. Cell Dev. Biol. 64 (April) (2017) 18–25. [84] L. Shi, X. Wang, Role of osteopontin in lung cancer evolution and heterogeneity, Semin. Cell Dev. Biol. 64 (April) (2017) 40–47. [85] M. Xu, D.C. Wang, X. Wang, Y. Zhang, Correlation between mucin biology and tumor heterogeneity in lung cancer, Semin. Cell Dev. Biol. 64 (April) (2017) 73–78. [86] L. Wang, B. Zhu, M. Zhang, X. Wang, Roles of immune microenvironment heterogeneity in therapy-associated biomarkers in lung cancer, Semin. Cell Dev. Biol. 64 (April) (2017) 90–97. [87] D. Wu, L. Zhuo, X. Wang, Metabolic reprogramming of carcinoma-associated fibroblasts and its impact on metabolic heterogeneity of tumors, Semin. Cell Dev. Biol. 64 (April) (2017) 125–131.

Please cite this article in press as: D.C. Wang, et al., A tour of 3D genome with a focus on CTCF, Semin Cell Dev Biol (2018), https://doi.org/10.1016/j.semcdb.2018.07.020