How enhancers activate their distal target promoters remains incompletely understood. Here we dissect how CTCF-mediated loops facilitate and restrict such regulatory interactions. Using an allelic series of mouse mutants, we show that CTCF is neither required for the interaction of the Sox2 gene with distal enhancers, nor for its expression. Insertion of various combinations of CTCF motifs, between Sox2 and its distal enhancers, generated boundaries with varying degrees of insulation that directly correlated with reduced transcriptional output. However, in both epiblast and neural tissues, enhancer contacts and transcriptional induction could not be fully abolished, and insertions failed to disrupt implantation and neurogenesis. In contrast, Sox2 expression was undetectable in the anterior foregut of mutants carrying the strongest boundaries, and these animals fully phenocopied loss of SOX2 in this tissue. We propose that enhancer clusters with a high density of regulatory activity can better overcome physical barriers to maintain faithful gene expression and phenotypic robustness.
This is a preview of subscription content, access via your institution
Access Nature and 54 other Nature Portfolio journals
Get Nature+, our best-value online-access subscription
$29.99 per month
cancel any time
Subscribe to this journal
Receive 12 print issues and online access
$189.00 per year
only $15.75 per issue
Rent or buy this article
Get just this article for as long as you need it
Prices may be subject to local taxes which are calculated during checkout
A list of publicly available data used in this study can be found in Supplementary Table 2, which includes data from refs. 22,94,106,107,108,109,110,111. All datasets were shown using processed files as available except for files mapped to mm9, which were converted to mm10 using CrossMap112. FASTQ and processed CHi-C and RAD21 E11.5 head ChIP-seq data can be found in GEO under accession number GSE190359 at ncbi.nlm.nih.gov/geo. CHi-C data can also be easily navigated at resgen.io/pedrorocha/sox2/views/.
Long, H. K., Prescott, S. L. & Wysocka, J. Ever-changing landscapes: transcriptional enhancers in development and evolution. Cell 167, 1170–1187 (2016).
de Laat, W. & Duboule, D. Topology of mammalian developmental enhancers and their regulatory landscapes. Nature 502, 499–506 (2013).
Schuijers, J. et al. Transcriptional dysregulation of MYC reveals common enhancer-docking mechanism. Cell Rep. 23, 349–360 (2018).
Kubo, N. et al. Promoter-proximal CTCF binding promotes distal enhancer-dependent gene activation. Nat. Struct. Mol. Biol. 28, 152–161 (2021).
Tang, Z. et al. CTCF-mediated human 3D genome architecture reveals chromatin topology for transcription. Cell 163, 1611–1627 (2015).
Vian, L. et al. The energetics and physiological impact of cohesin extrusion. Cell 173, 1165–1178 (2018).
Kane, L. et al. Cohesin is required for long-range enhancer action at the Shh locus. Nat. Struct. Mol. Biol. 29, 891–897 (2022).
Nora, E. P. et al. Targeted degradation of CTCF decouples local insulation of chromosome domains from genomic compartmentalization. Cell 169, 930–944 (2017).
Schwarzer, W. et al. Two independent modes of chromatin organization revealed by cohesin removal. Nature 551, 51–56 (2017).
Rao, S. S. P. et al. Cohesin loss eliminates all loop domains. Cell 171, 305–320 (2017).
Fudenberg, G. et al. Formation of chromosomal domains by loop extrusion. Cell Rep. 15, 2038–2049 (2016).
Li, Y. et al. The structural basis for cohesin-CTCF-anchored loops. Nature 578, 472–476 (2020).
Deng, W. et al. Controlling long-range genomic interactions at a native locus by targeted tethering of a looping factor. Cell 149, 1233–1244 (2012).
Weintraub, A. S. et al. YY1 Is a structural regulator of enhancer–promoter loops. Cell 171, 1573–1588 (2017).
Richter, W. F., Nayak, S., Iwasa, J. & Taatjes, D. J. The mediator complex as a master regulator of transcription by RNA polymerase II. Nat. Rev. Mol. Cell Biol. 23, 732–749 (2022).
Pombo, A. & Dillon, N. Three-dimensional genome architecture: players and mechanisms. Nat. Rev. Mol. Cell Biol. 16, 245–257 (2015).
Oudelaar, A. M. & Higgs, D. R. The relationship between genome structure and function. Nat. Rev. Genet. 22, 154–168 (2021).
Furlong, E. E. M. & Levine, M. Developmental enhancers and chromosome topology. Science 361, 1341–1345 (2018).
Schoenfelder, S. & Fraser, P. Long-range enhancer–promoter contacts in gene expression control. Nat. Rev. Genet. 20, 437–455 (2019).
Zhang, S., Übelmesser, N., Barbieri, M. & Papantonis, A. Enhancer–promoter contact formation requires RNAPII and antagonizes loop extrusion. Preprint at bioRxiv https://doi.org/10.1101/2022.07.04.498738 (2022).
Sexton, T. et al. Three-dimensional folding and functional organization principles of the Drosophila genome. Cell 148, 458–472 (2012).
Bonev, B. et al. Multiscale 3D genome rewiring during mouse neural development. Cell 171, 557–572 (2017).
Rao, S. S. et al. A 3D map of the human genome at kilobase resolution reveals principles of chromatin looping. Cell 159, 1665–1680 (2014).
Nora, E. P. et al. Spatial partitioning of the regulatory landscape of the X-inactivation centre. Nature 485, 381–385 (2012).
Dixon, J. R. et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485, 376–380 (2012).
Lupianez, D. G. et al. Disruptions of topological chromatin domains cause pathogenic rewiring of gene–enhancer interactions. Cell 161, 1012–1025 (2015).
Narendra, V. et al. CTCF establishes discrete functional chromatin domains at the Hox clusters during differentiation. Science 347, 1017–1021 (2015).
Kraft, K. et al. Serial genomic inversions induce tissue-specific architectural stripes, gene misexpression and congenital malformations. Nat. Cell Biol. 21, 305–310 (2019).
Engel, N., Raval, A. K., Thorvaldsen, J. L. & Bartolomei, S. M. Three-dimensional conformation at the H19/Igf2 locus supports a model of enhancer tracking. Hum. Mol. Genet. 17, 3021–3029 (2008).
Despang, A. et al. Functional dissection of the Sox9–Kcnj2 locus identifies nonessential and instructive roles of TAD architecture. Nat. Genet. 51, 1263–1271 (2019).
Ghavi-Helm, Y. et al. Highly rearranged chromosomes reveal uncoupling between genome topology and gene expression. Nat. Genet. 51, 1272–1282 (2019).
Rodriguez-Carballo, E. et al. Chromatin topology and the timing of enhancer function at the HoxD locus. Proc. Natl Acad. Sci. USA 117, 31231–31241 (2020).
Williamson, I. et al. Developmentally regulated Shh expression is robust to TAD perturbations. Development 146, dev179523 (2019).
Amandio, A. R. et al. Sequential in cis mutagenesis in vivo reveals various functions for CTCF sites at the mouse HoxD cluster. Genes Dev. 35, 1490–1509 (2021).
Paliou, C. et al. Preformed chromatin topology assists transcriptional robustness of Shh during limb development. Proc. Natl Acad. Sci. USA 116, 12390–12399 (2019).
Misteli, T. The self-organizing genome: principles of genome architecture and function. Cell 183, 28–45 (2020).
Beagan, J. A. & Phillips-Cremins, J. E. On the existence and functionality of topologically associating domains. Nat. Genet. 52, 8–16 (2020).
Mirny, L. & Dekker, J. Mechanisms of chromosome folding and nuclear organization: their interplay and open questions. Cold Spring Harb. Perspect. Biol. 14, a040147 (2022).
Avilion, A. A. et al. Multipotent cell lineages in early mouse development depend on SOX2 function. Genes Dev. 17, 126–140 (2003).
Li, Y. et al. CRISPR reveals a distal super-enhancer required for Sox2 expression in mouse embryonic stem cells. PLoS ONE 9, e114485 (2014).
Zhou, H. Y. et al. A Sox2 distal enhancer cluster regulates embryonic stem cell differentiation potential. Genes Dev. 28, 2699–2711 (2014).
Taylor, T. et al. Transcriptional regulation and chromatin architecture maintenance are decoupled functions at the Sox2 locus. Genes Dev. 36, 699–717 (2022).
Brosh, R. et al. Synthetic regulatory genomics uncovers enhancer context dependence at the Sox2 locus. Preprint at bioRxiv https://doi.org/10.1101/2022.06.20.495832 (2022).
Saiz, N., Kang, M., Schrode, N., Lou, X. & Hadjantonakis, A. K. Quantitative analysis of protein expression to study lineage specification in mouse preimplantation embryos. J. Vis. Exp. 22, 53654 (2016).
Chazaud, C., Yamanaka, Y., Pawson, T. & Rossant, J. Early lineage segregation between epiblast and primitive endoderm in mouse blastocysts through the Grb2-MAPK pathway. Dev. Cell 10, 615–624 (2006).
Wicklow, E. et al. HIPPO pathway members restrict SOX2 to the inner cell mass where it promotes ICM fates in the mouse blastocyst. PLoS Genet. 10, e1004618 (2014).
Anania, C. et al. In vivo dissection of a clustered-CTCF domain boundary reveals developmental principles of regulatory insulation. Nat. Genet. 54, 1026–1036 (2022).
Chang, L.-H. et al. A complex CTCF binding code defines TAD boundary structure and function. Preprint at bioRxiv https://doi.org/10.1101/2021.04.15.440007 (2021).
Ferri, A. L. et al. Sox2 deficiency causes neurodegeneration and impaired neurogenesis in the adult mouse brain. Development 131, 3805–3819 (2004).
Li, P. Y., Li, S. Q., Gao, S. G. & Dong, D. Y. CRISPR/Cas9-mediated gene editing on Sox2ot promoter leads to its truncated expression and does not influence neural tube closure and embryonic development in mice. Biochem. Biophys. Res. Commun. 573, 107–111 (2021).
Denker, A. & de Laat, W. The second decade of 3C technologies: detailed insights into nuclear organization. Genes Dev. 30, 1357–1382 (2016).
Redolfi, J. et al. DamC reveals principles of chromatin folding in vivo without crosslinking and ligation. Nat. Struct. Mol. Biol. 26, 471–480 (2019).
Vos, E. S. M. et al. Interplay between CTCF boundaries and a super enhancer controls cohesin extrusion trajectories and gene expression. Mol. Cell 81, 3082–3095 (2021).
Benabdallah, N. S. et al. Decreased enhancer–promoter proximity accompanying enhancer activation. Mol. Cell 76, 473–484 (2019).
Alexander, J. M. et al. Live-cell imaging reveals enhancer-dependent Sox2 transcription in the absence of enhancer proximity. eLife 8, e41769 (2019).
Fulco, C. P. et al. Activity-by-contact model of enhancer-promoter regulation from thousands of CRISPR perturbations. Nat. Genet. 51, 1664–1669 (2019).
Favaro, R. et al. Hippocampal development and neural stem cell maintenance require Sox2-dependent regulation of Shh. Nat. Neurosci. 12, 1248–1256 (2009).
Beagan, J. A. et al. YY1 and CTCF orchestrate a 3D chromatin looping switch during early neural lineage commitment. Genome Res. 27, 1139–1152 (2017).
Uchikawa, M. & Kondoh, H. in Sox2 (eds. Kondoh, H. & Lovell-Badge, R.) 107–129 (Academic Press, 2016).
Edwards, N. A. et al. Developmental basis of trachea-esophageal birth defects. Dev. Biol. 477, 85–97 (2021).
Que, J. et al. Multiple dose-dependent roles for Sox2 in the patterning and differentiation of anterior foregut endoderm. Development 134, 2521–2531 (2007).
Teramoto, M. et al. The absence of SOX2 in the anterior foregut alters the esophagus into trachea and bronchi in both epithelial and mesenchymal components. Biol. Open 9, bio048728 (2020).
Zenteno, J. C., Perez-Cano, H. J. & Aguinaga, M. Anophthalmia-esophageal atresia syndrome caused by an SOX2 gene deletion in monozygotic twin brothers with markedly discordant phenotypes. Am. J. Med. Genet. A 140, 1899–1903 (2006).
Langer, L., Sulik, K. & Pevny, L. Cleft palate in a mouse model of SOX2 haploinsufficiency. Cleft Palate Craniofac. J. 51, 110–114 (2014).
Mandalos, N. et al. Sox2 acts as a rheostat of epithelial to mesenchymal transition during neural crest development. Front. Physiol. 5, 345 (2014).
Arnold, K. et al. Sox2+ adult stem and progenitor cells are important for tissue regeneration and survival of mice. Cell Stem Cell 9, 317–329 (2011).
Ibarra-Soria, X. et al. Defining murine organogenesis at single-cell resolution reveals a role for the leukotriene pathway in regulating blood progenitor formation. Nat. Cell Biol. 20, 127–134 (2018).
Dowen, J. M. et al. Control of cell identity genes occurs in insulated neighborhoods in mammalian chromosomes. Cell 159, 374–387 (2014).
Flavahan, W. A. et al. Insulator dysfunction and oncogene activation in IDH mutant gliomas. Nature 529, 110–114 (2016).
Laugsch, M. et al. Modeling the pathological long-range regulatory effects of human structural variation with patient-specific hiPSCs. Cell Stem Cell 24, 736–752 (2019).
van Bemmel, J. G. et al. The bipartite TAD organization of the X-inactivation center ensures opposing developmental regulation of Tsix and Xist. Nat. Genet. 51, 1024–1034 (2019).
Hanssen, L. L. P. et al. Tissue-specific CTCF-cohesin-mediated chromatin architecture delimits enhancer interactions and function in vivo. Nat. Cell Biol. 19, 952–961 (2017).
de Wit, E. et al. CTCF binding polarity determines chromatin looping. Mol. Cell 60, 676–684 (2015).
Aljahani, A. et al. Analysis of sub-kilobase chromatin topology reveals nano-scale regulatory interactions with variable dependence on cohesin and CTCF. Nat. Commun. 13, 2139 (2022).
Zhang, H. et al. CTCF and transcription influence chromatin structure re-configuration after mitosis. Nat. Commun. 12, 5157 (2021).
Hsieh, T. S. et al. Enhancer-promoter interactions and transcription are largely maintained upon acute loss of CTCF, cohesin, WAPL or YY1. Nat. Genet. 54, 1919–1932 (2022).
Zuin, J. et al. Nonlinear control of transcription through enhancer-promoter interactions. Nature 604, 571–577 (2022).
Symmons, O. et al. The Shh topological domain facilitates the action of remote enhancers by reducing the effects of genomic distances. Dev. Cell 39, 529–543 (2016).
Xiao, J. Y., Hafner, A. & Boettiger, A. N. How subtle changes in 3D structure can create large changes in transcription. eLife 10, e64320 (2021).
Rinzema, N. J. et al. Building regulatory landscapes reveals that an enhancer can recruit cohesin to create contact domains, engage CTCF sites and activate distant genes. Nat. Struct. Mol. Biol. 29, 563–574 (2022).
Huang, H. et al. CTCF mediates dosage- and sequence-context-dependent transcriptional insulation by forming local chromatin domains. Nat. Genet. 53, 1064–1074 (2021).
Galupa, R. et al. A conserved noncoding locus regulates random monoallelic Xist expression across a topological boundary. Mol. Cell 77, 352–367 (2020).
Pinto, P. B. et al. Enhancer-promoter interactions form independently of genomic distance and are functional across TAD boundaries. Preprint at bioRxiv https://doi.org/10.1101/2022.08.29.505755 (2022).
Schmidt, D. et al. Waves of retrotransposon expansion remodel genome organization and CTCF binding in multiple mammalian lineages. Cell 148, 335–348 (2012).
Sundaram, V. et al. Widespread contribution of transposable elements to the innovation of gene regulatory networks. Genome Res. 24, 1963–1976 (2014).
Vietri Rudan, M. et al. Comparative Hi-C reveals that CTCF underlies evolution of chromosomal domain architecture. Cell Rep. 10, 1297–1309 (2015).
Bourque, G. et al. Evolution of the mammalian transcription factor binding repertoire via transposable elements. Genome Res. 18, 1752–1762 (2008).
Lunyak, V. V. et al. Developmentally regulated activation of a SINE B2 repeat as a domain boundary in organogenesis. Science 317, 248–251 (2007).
Gu, H. et al. Fine-mapping of nuclear compartments using ultra-deep Hi-C shows that active promoter and enhancer elements localize in the active A compartment even when adjacent sequences do not. Preprint at bioRxiv https://doi.org/10.1101/2021.10.03.462599 (2021).
Goel, V. Y., Huseyin, M. K. & Hansen, A. S. Region Capture Micro-C reveals coalescence of enhancers and promoters into nested microcompartments. Preprint at bioRxiv https://doi.org/10.1101/2022.07.12.499637 (2022).
Rowley, M. J. et al. Evolutionarily conserved principles predict 3D chromatin organization. Mol. Cell 67, 837–852 (2017).
Hnisz, D., Shrinivas, K., Young, R. A., Chakraborty, A. K. & Sharp, P. A. A phase separation model for transcriptional control. Cell 169, 13–23 (2017).
Oudelaar, A. M. et al. Single-allele chromatin interactions identify regulatory hubs in dynamic compartmentalized domains. Nat. Genet. 50, 1744–1751 (2018).
Thompson, J. J. et al. Extensive co-binding and rapid redistribution of NANOG and GATA6 during emergence of divergent lineages. Nat. Commun. 13, 4257 (2022).
Chari, R., Yeo, N. C., Chavez, A. & Church, G. M. sgRNA Scorer 2.0: a species-independent model to predict CRISPR/Cas9 activity. ACS Synth. Biol. 6, 902-904 (2017).
Gooden, A. A., Evans, C. N., Sheets, T. P., Clapp, M. E. & Chari, R. dbGuide: a database of functionally validated guide RNAs for genome editing in human and mouse cells. Nucleic Acids Res. 49, D871–D876 (2021).
D’Astolfo, D. S. et al. Efficient intracellular delivery of native proteins. Cell 161, 674–690 (2015).
Czechanski, A. et al. Derivation and characterization of mouse embryonic stem cells from permissive and nonpermissive strains. Nat. Protoc. 9, 559–574 (2014).
Mali, P. et al. RNA-guided human genome engineering via Cas9. Science 339, 823–826 (2013).
Moorthy, S. D. et al. Enhancers and super-enhancers have an equivalent regulatory role in embryonic stem cells through regulation of single or multiple genes. Genome Res. 27, 246–258 (2017).
Ding, Q. et al. Enhanced efficiency of human pluripotent stem cell genome editing through replacing TALENs with CRISPRs. Cell Stem Cell 12, 393–394 (2013).
Zalc, A. et al. Reactivation of the pluripotency program precedes formation of the cranial neural crest. Science 371, eabb4776 (2017).
Servant, N. et al. HiC-Pro: an optimized and flexible pipeline for Hi-C data processing. Genome Biol. 16, 259 (2015).
Kerpedjiev, P. et al. HiGlass: web-based visual exploration and analysis of genome interaction maps. Genome Biol. 19, 125 (2018).
Kruse, K., Hug, C. B. & Vaquerizas, J. M. FAN-C: a feature-rich framework for the analysis and visualisation of chromosome conformation capture data. Genome Biol. 21, 303 (2020).
Hansen, A. S., Pustova, I., Cattoglio, C., Tjian, R. & Darzacq, X. CTCF and cohesin regulate chromatin loop stability with distinct dynamics. eLife 6, e25776 (2017).
Kagey, M. H. et al. Mediator and cohesin connect gene expression and chromatin architecture. Nature 467, 430–435 (2010).
Whyte, W. A. et al. Master transcription factors and mediator establish super-enhancers at key cell identity genes. Cell 153, 307–319 (2013).
Xiang, Y. et al. Epigenomic analysis of gastrulation identifies a unique chromatin state for primed pluripotency. Nat. Genet. 52, 95–105 (2020).
Gorkin, D. U. et al. An atlas of dynamic chromatin landscapes in mouse fetal development. Nature 583, 744–751 (2020).
Li, J. et al. Accurate annotation of accessible chromatin in mouse and human primordial germ cells. Cell Res. 28, 1077–1089 (2018).
Zhao, H. et al. CrossMap: a versatile tool for coordinate conversion between genome assemblies. Bioinformatics 30, 1006–1007 (2014).
We would like to thank all members of the Unit on Genome Structure and Regulation for comments and discussions on this project and manuscript as well as K. Pfeifer, T. Macfarlan and J. Kassis. We thank H. Schrewe for tips on AFG isolation and N. Saiz for blastocyst IF and image analysis. We thank NICHD’s molecular genetics core, specifically S. Coon, T. Li and J. Iben. This work uses the computational resources of the NIH HPC Biowulf cluster (hpc.nih.gov). We thank L. Price for cell sorting experiments. We thank the mouse core of NICHD, specifically J. Yimdjo, V. Biggs and A. Grinberg. We also thank the NCI’s molecular histopathology core, especially T. Morgan, J. Mata and B. Karim. This work was funded by NIH intramural project HD008975 (to P.P.R.), HD008986 (to R.K.D.), HD008962 (to T.J.P.) and the Canadian Institutes of Health Research FRN 153186 (to J.A.M.). This project has also been funded in part with Federal funds from the National Cancer Institute, National Institutes of Health, under contract HHSN261201500003I. The content of this publication does not necessarily reflect the views or policies of the Department of Health and Human Services nor does mention of trade names, commercial products or organizations imply endorsement by the US Government.
The authors declare no competing interests.
Peer review information
Nature Genetics thanks Bing Ren and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended Data Fig. 1 SCR is required for Sox2 expression.
a Sox2 and SCR activity during early mouse development as assessed by enrichment of H3K27ac and ATAC-seq. SCR is active in pluripotent epiblast cells, and in germ cells that regain a pluripotent status. H3K27ac CUT&RUN in mES cells cultured in epiblast-like conditions shows enrichment at both Sox2 and SCR but not in cells differentiated into primitive endoderm (PrE). No H3K27ac is detected at Sox2 or SCR in the visceral endoderm of post-implantation embryos and SCR activity is diminished in epiblast cells. Despite strong enrichment at the Sox2 gene body in the neural tube and forebrain of E11.5 embryos, SCR is no longer active. Instead, other proximal regions (black arrow) show increased activity. ATAC-seq data shows that both male and female primordial germ cells, but not the surrounding soma, are enriched in accessible chromatin at SCR. b Regions targeted for deletion in the SCR∆ line and gRNAs used. Browser shot shows enrichment of NANOG, SOX2, OCT4 and H3K27ac over Sox2 and SCR in ES cells. Nucleotides in red represent the protospacer sequence of gRNAs used for injection. Underlined nucleotides highlight location of the Cas9 PAM in the mouse genome. Regions targeted in previous studies are show also with black boxes. Arrows on both sides of the deletion highlight the ligation junction detected in the mouse used as founder as determined by Sanger sequencing. c IF of E4.5 blastocysts stained with antibodies targeting GATA6, NANOG and SOX2. In the quantification plots, each dot represents a cell and to allow comparison across three different litters, the intensity of each cell was normalized by the cell with highest intensity in heterozygous embryos. Number of embryos analyzed for each genotype is shown below the plot. Boxplots show minimum, maximum, median, first, and third quartiles. A Wilcoxon two-sided test was performed to assess statistical significance. Scale bar represents 10 µm.
Extended Data Fig. 2 SCR activates Sox2 independently of CTCF.
a Scheme depicting targeting strategy for generation of the two CTCF deletion lines. Browser shot shows enrichment of CTCF, NANOG, SOX2, OCT4 and H3K27ac over Sox2 and SCR. A NANOG/SOX2/OCT4 peak showing high enrichment was deleted in the CTCF∆(C2-C4) but this region has been previously shown to not be required for mouse development. Browser shots with zoomed-in views of CTCF enrichment at Sox2 and SCR show precise location of gRNAs used in zygotic injections. CTCF peak nearest the most centromeric gRNA used in the CTCF∆(C2-C4) targeting does not contain a significant CTCF motif according to FIMO. This peak was nonetheless deleted in the CTCF∆(C2-C4) line. Nucleotides in red represent the protospacer sequence of gRNAs used for injection. Underlined nucleotides highlight location of the Cas9 PAM in the mouse genome. Arrows on both sides of the deletion highlight ligation junction detected in the mouse used as founder as determined by Sanger sequencing. For the CTCF∆(C5) scheme, purple line highlights location of the CTCF motif at SCR. The central sequence of the repair template is shown, with parts of the protospacer sequence used in the gRNA shown in red, and the restriction enzyme target that replaced the CTCF motif in blue. Full sequence of homology arms on both sides is omitted. b Differential CHi-C interaction frequency heatmaps. Red signal represents interactions occurring at higher frequency in mutant cell lines compared to control and blue shows interactions of lower frequency. Dotted lines represent the Sox2-SCR domain as detected in WT control cells.
Extended Data Fig. 3 SCR induces Sox2 independently of CTCF.
a Virtual 4C plots using the Sox2 and SCR viewpoints at DpnII-fragment resolution. Plots in main figure show overlapping 5 kb windows. These plots can be used to also visualize the maintenance of Sox2-SCR interactions. Region surrounding viewpoint was removed from analysis. Dotted lines highlight SCR in the Sox2 viewpoint (left), and Sox2 in the SCR viewpoint (right). Virtual 4C signal is shown as average of the 2 replicates at the fragment level. b Deletion of CTCF motifs on the centromeric and telomeric end of the Sox2-SCR domain does not affect interactions and Sox2 expression. Left-qPCR analysis of Sox2 expression in ES cells was done using the ∆∆CT method and Gapdh as a reference. Sox2 expression was calculated by comparing it to the median WT clones. Each dot represents a different ES cell clone. The number of independent cell lines(n) analyzed of each genotype was 2. Boxplots show minimum, maximum, median, first, and third quartiles. A Wilcoxon two-sided test was performed to assess statistical significance. Right-CHi-C 1D interaction frequency heatmap in homozygotic CTCF∆(C2-C4) + (C5) mES cells compared to WT.
Extended Data Fig. 4 SCR can activate Sox2 across CTCF-mediated insulation.
a Top left scheme depicts targeting strategy for generation of the transgenic lines carrying insertion of CTCF motifs. Arcs represent CTCF-mediated loops generated in each insertion line as predicted by the loop-extrusion model. We assumed that the sites where cassettes were targeted did not contain regulatory elements based on conservation and analysis of ENCODE datasets. Top right browser shows CTCF ChIP-seq enrichment and insertion sites of CTCF transgenes. Left bottom panel shows the targeting of the C57Bl6 genome to generate CTCFi3×− mice. Right bottom panel shows targeting to generate CTCFi3×+, CTCFi3×−;3×+, and CTCFi18×+ mice. Nucleotides shown in red represent the protospacer sequence of gRNAs used for injection while underlined nucleotides highlight location of the Cas9 PAM in the mouse mm10 genome. gRNA mm10 coordinates are shown in red. The central sequence of the repair templates is shown, with parts of the protospacer sequence used in the gRNA shown in red and mutated PAM nucleotides underlined. The complete sequence of homology arms on both sides is omitted. Each colored rectangle represents a different region from the mouse genome containing a CTCF motif and adjacent regions. Complete sequences of the CTCF motifs and adjacent regions are shown in different color with the central CTCF motif in blue representing targeting to negative strand in the CTCFi3×− line. The first and last two nucleotides of each of the three CTCF regions are shown in the schema of the repair template. The same three CTCF-carrying regions were used in CTCFi3×− and CTCFi3×+ lines but targeted to different strands and locations. Therefore, the central region of repair template is the same but in different strands and containing different repair templates. Retargeting of the CTCFi3×+ transgene on a homozygous CTCFi3×− background generated the CTCFi3×−;3×+ line. The CTCFi18×+ line was obtained as a consequence of the CTCFi3×+ injection because of concatemerization of the repair template. The resulting allele is shown in the bottom right. b Differential CHi-C interaction frequency heatmap. Red signal represents interactions occurring at higher frequency in mutant cell lines compared to control. Dotted lines represent insertion sites of CTCF transgenes.
Extended Data Fig. 5 SCR can interact with Sox2 across CTCF-mediated insulation.
a Virtual 4C plot using Sox2 and SCR viewpoints. Region surrounding viewpoint was removed from analysis. Dotted lines highlight SCR in the Sox2 viewpoint (top), and Sox2 in the SCR viewpoint (bottom). Virtual 4C signal is shown as average of the 2 replicates of each genotype in 5 kb overlapping windows. b Virtual 4C plot using Sox2 and SCR viewpoints at DpnII-fragment resolution. This same signal at 5 kb overlapping bins is shown in A and in the main figure. This representation highlights how, despite massively reduced, Sox2 can interact with SCR in all lines generated. Region surrounding viewpoint was removed from analysis. Dotted lines highlight SCR in the Sox2 viewpoint (left), and Sox2 in the SCR viewpoint (right). Virtual 4C signal is shown as average of the 2 replicates of each genotype at DpnII fragment resolution. c Results from ABC model using WT H3K27ac and CHi-C data from each of our mutants. The lower score in CTCF∆(C2-C4) mutants is likely related to the 5 kb windows used for analysis, which cause a very strong artificial reduction in CHi-C signal on these cells that have an 8 kb deletion very near the Sox2 promoter.
Extended Data Fig. 6 DNE also induces Sox2 across CTCF-mediated insulation.
a Interaction frequency heatmap determined by CHi-C in heads of E11.5 embryos. Insets on the right show 2D interaction heatmaps highlighting interactions between regions surrounding Sox2 and DNE. CTCF data shown under CHi-C heatmaps from heads is derived from in vitro differentiated neural progenitor cells. Rectangles represent the Sox2-DNE interaction, arrowheads represent loops with CTCF downstream of DNE established by transgene insertions, black arrow represent loops with CTCF upstream of Sox2 established by transgene insertions. b Insulation scores for 5 kb windows in this region are shown where lower levels represent higher insulation. CTCFi18×+ shows the strongest insulation score while CTCFi3×−;3×+ displays the largest ectopic boundary. c Differential CHi-C interaction frequency heatmap. Red signal represents interactions occurring at higher frequency in mutant cell lines compared to control. Black arrow highlights formation of a highly interacting domain containing Sox2 and the proximal neural enhancers.
Extended Data Fig. 7 DNE can activate Sox2 across CTCF-mediated insulation.
a Virtual 4C plot using Sox2 and DNE viewpoints with signal plotted at each DpnII fragment. This same signal at 5 kb overlapping bins is shown in the main figure. This representation highlights how, despite massively reduced, Sox2 can interact with DNE in all lines assessed. Signal is shown as average of the 2 replicates of each genotype. Region surrounding viewpoint was removed from analysis. Dotted lines highlight DNE in the Sox2 viewpoint (left), and Sox2 in the DNE viewpoint (right). Virtual 4C signal is shown as average of the 2 replicates of each genotype at DpnII fragment resolution. b IF of E11.5 embryos stained with an antibody targeting SOX2. Scale bar represents 500 µm. 2 embryos of each genotype were stained and imaged.
Extended Data Fig. 8 Developmental defects seen in homozygous embryos with modifications of the Sox2 locus.
a Transverse section of E13.5 CTCFi3×−;3×+ wt and homozygote littermates at heart level. Es-esophagus, Tr-trachea. 3 embryos from each genotype were sectioned, stained and imaged. Scale bar represents 100 µm b Genotyping at of living animals at weaning, P0 and E18.5 for the indicated strains. P values were calculated using a two-tailed chi-squared test. The viability of homozygous CTCFi3×− and CTCFi3×+ animals provides strong evidence that insertion of CTCF cassettes did not disrupt regulatory elements and that the phenotypes seen in CTCFi3×−;3×+ and CTCFi18×+ homozygotes are caused by perturbations to the chromatin structure of the Sox2 locus. In agreement with our observation that 1 of 9 post-implantation E6.5 mutant embryos initiated gastrulation (Fig. 1c), we recovered a few SCR∆ homozygotes at weaning but at a highly reduced frequency compared to the expected ratio. This could be explained by SCR losing activity following implantation, suggesting that embryos that successfully implant despite SCR deletion can complete development. In line with absence of SOX2 expression in the AFG of E9.5 embryos, we only recovered 3 CTCFi18×+homozygous pups alive at P0 (out of 15 expected) and none at weaning. As the three pups found at P0 were observed during delivery we speculate that they would perish within a few hours as all other analyzed pups of this line. c Frontal sections of E18.5 CTCFi3×−;3×+ wt and homozygote littermates. Asterisk highlights cleft palate defect. 3 embryos from each genotype were sectioned, stained and imaged. Scale bar represents 500 µm. d Plot of normalized Sox2 expression in single cells of WT E8.25 embryos. 16 biologically independent embryos (n) were analyzed. Boxplots show minimum, maximum, median, first, and third quartiles. Plot was modified from marionilab.cruk.cam.ac.uk/organogenesis/.
Supplementary Table 1
List of antibodies used, sequence of PCR oligos, gRNAs, repair templates and deletion junctions.
Supplementary Table 2
Statistics on sequencing data, list of public datasets used, and genomic location of CHi-C probes used.
Raw nanopore sequencing files of founder mice.
Rights and permissions
Springer Nature or its licensor (e.g. a society or other partner) holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law.
About this article
Cite this article
Chakraborty, S., Kopitchinski, N., Zuo, Z. et al. Enhancer–promoter interactions can bypass CTCF-mediated boundaries and contribute to phenotypic robustness. Nat Genet 55, 280–290 (2023). https://doi.org/10.1038/s41588-022-01295-6