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Tissue-specific CTCF–cohesin-mediated chromatin architecture delimits enhancer interactions and function in vivo

Nature Cell Biology volume 19, pages 952–961 (2017)Cite this article

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Abstract

The genome is organized via CTCF–cohesin-binding sites, which partition chromosomes into 1–5 megabase (Mb) topologically associated domains (TADs), and further into smaller sub-domains (sub-TADs). Here we examined in vivo an ∼80 kb sub-TAD, containing the mouse α-globin gene cluster, lying within a ∼1 Mb TAD. We find that the sub-TAD is flanked by predominantly convergent CTCF–cohesin sites that are ubiquitously bound by CTCF but only interact during erythropoiesis, defining a self-interacting erythroid compartment. Whereas the α-globin regulatory elements normally act solely on promoters downstream of the enhancers, removal of a conserved upstream CTCF–cohesin boundary extends the sub-TAD to adjacent upstream CTCF–cohesin-binding sites. The α-globin enhancers now interact with the flanking chromatin, upregulating expression of genes within this extended sub-TAD. Rather than acting solely as a barrier to chromatin modification, CTCF–cohesin boundaries in this sub-TAD delimit the region of chromatin to which enhancers have access and within which they interact with receptive promoters.

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Figure 1: Regulation of the α-globin cluster in mouse ES and primary erythroid cells.
Figure 2: Differential interactions of α-globin regulatory elements between mouse ES and primary erythroid cells.
Figure 3: Disruption of CTCF binding motifs results in loss of CTCF binding at the α-globin locus in primary erythroid cells derived from mutant mice.
Figure 4: Differential interactions of α-globin regulatory regions and flanking genes between WT and D3839 primary erythroid cells.
Figure 5: Effects of individual and combined CTCF binding site deletions near the α-globin enhancers on local gene expression in primary erythroid cells.
Figure 6: Effects of combined deletion of HS-38 and HS-39 on the local chromatin state in primary erythroid cells.

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Acknowledgements

The authors would like to thank J. Telenius for her help with bioinformatic analysis of the data and D. Hay, R. Gibbons and A. King for critically reading the manuscript. L.L.P.H. and A.M.O. would like to thank the Wellcome Trust for funding (Chromosome and Developmental Biology PhD Programme, grant code 099684/Z/12/Z; and Wellcome Trust Genomic Medicine and Statistics PhD Programme, grant code 083323/Z/07/Z respectively). The work was further supported by the Wellcome Trust core award 203141/Z/16/Z and the Medical Research Council (MRC Core Funding and Centenary Award reference 4050189188).

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Author notes

  1. Mira T. Kassouf and A. Marieke Oudelaar: These authors contributed equally to this work.

  2. Benjamin Davies and Douglas R. Higgs: These authors jointly supervised this work.

Authors and Affiliations

  1. MRC Molecular Haematology Unit, Weatherall Institute of Molecular Medicine, Oxford, OX3 9DS, UK

    Lars L. P. Hanssen, Mira T. Kassouf, A. Marieke Oudelaar, Damien J. Downes, Matthew Gosden, Jacqueline A. Sharpe, Jacqueline A. Sloane-Stanley, Jim R. Hughes & Douglas R. Higgs

  2. The Wellcome Trust Centre for Human Genetics, Roosevelt Drive, University of Oxford, Oxford, OX3 7BN, UK

    Lars L. P. Hanssen, Daniel Biggs, Chris Preece & Benjamin Davies

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Contributions

L.L.P.H. planned and carried out experiments, analysed the data, carried out bioinformatic analysis, and wrote the manuscript. M.T.K. coordinated and advised on the project and revised the manuscript. A.M.O. carried out Capture-C experiments and revised the manuscript. M.T.K. and A.M.O. contributed equally to this work. D.B. carried out cell culture and mouse experiments. C.P. carried out mouse microinjection experiments. D.J.D. and M.G. carried out ATAC-seq experiments. J.A.S. and J.A.S.-S. carried out mouse maintenance and haematology experiments. J.R.H. coordinated the project and gave technical and bioinformatics advice throughout the project. B.D. planned and coordinated transgenic mouse experiments and wrote the manuscript. D.R.H. conceived, planned and coordinated the project and wrote the manuscript. B.D. and D.R.H. share senior authorship.

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Correspondence to Douglas R. Higgs.

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Integrated supplementary information

Supplementary Figure 1 De novo analysis of the CTCF binding motifs and footprints in mouse erythroid cells.

(a) CTCF consensus binding motif in forward and reverse orientation resulting from MEME motif analysis of CTCF ChIP-seq (2 independent experiments in which 2 animals were analysed in total) in erythroid cells. Core (C), upstream (U), and downstream (D) sequence elements are identified with preferential spacing to the core motif (spacer, red bar histograms). (b) Plots of the average DNaseI footprints of C, UC, and UCD motif containing CTCF binding sites in forward orientation (top panel) and reverse orientation (lower panel). Upper (red, +) and lower (blue, -) strand specific DNaseI cleavage signals are shown. Footprints are averaged over the total number of sites in each category (indicated between parentheses). (c) Normalised CTCF ChIP-seq (RPKM, 2 independent experiments in which 2 animals were analysed in total) annotated with the CTCF site names and orientation at the α-globin locus in erythroid cells. Gene annotation is Refseq. DNaseI footprints and top CTCF binding motif hit for each of the CTCF binding sites. Motif P-values are shown (as explained in Fig. 1c). Orientation is indicated by the colour of the arrow over the core motif (C); forward (red) or reverse (blue). Upstream motif (U) is shown in green and downstream motif (D) in yellow.

Supplementary Figure 2 Genome editing strategies for disruption of CTCF binding at the α-globin locus.

Schematic overview of the mutagenesis method (CRISPR-Cas9 and/or TALEN) and the resulting mutations induced at HS-38 (D38), HS-39 (D39), HS-38 and HS-39 (D3839), and HS-29 (D29). DNaseI footprints are annotated with matches to the C, U, and D CTCF motif (coloured bar under C, red = forward, blue = reverse). Where homology directed repair was used to replace part of the CTCF core motif with a restriction site, the single-stranded oligodeoxynucleotides (ssODNs) used as repair templates are shown.

Supplementary Figure 3 Differential interactions of α-globin regulatory regions and flanking genes between wild-type and D29 primary erythroid cells.

Capture-C interaction data for the indicated viewpoints (red vertical bars) in wild-type (WT) and CTCF mutant D29 (deletion of HS-29) primary erythroid cells is shown as described in Fig. 2a. Data represent 3 independent experiments in which 3 animals were analysed in total. Shaded grey bar indicates the mutated CTCF site (HS-29). Also shown are normalised CTCF ChIP-seq (RPKM) for both WT and D29 erythroid cells and ATAC-seq (RPKM) for WT erythroid cells, all merged from 2 independent experiments in which 2 animals were analysed. Gene annotation is Refseq.

Supplementary Figure 4 Effects of the deletion of HS-29 on local gene expression and chromatin state in primary erythroid cells.

(a) MA plot for RNA-seq data derived from WT and D29 erythroid cells. Data represent n = 3 independent experiments in which 3 animals were analysed in total. Mean RNA abundance is plotted on the x-axis and enrichment is plotted on the y-axis. No significant changes in local gene expression were detected as shown on the plot by the genes highlighted in blue: Snrnp25, Rhbdf1, Mpg. Controls are shown in different colours: Mitoferrin-1 (Slc25a37, green), a highly expressed erythroid gene; Sh3pxd2b and Il9r (red), repressed in erythroid; Nprl3 (yellow), a housekeeping gene within the α-globin locus that is unaffected by HS-29 deletion. The five most significant outliers were investigated and only one gene was located in cis to α-globin (Irgm2, >25 Mb removed), making it unlikely that these expression changes represent direct effects from HS-29 deletion. Significance was tested with a Wald test (DEseq2, n = 3 independent experiments in which 3 animals were analysed). (b) Normalised ATAC-Seq and ChIP-seq read-densities (RPKM; 2 independent experiments in which 2 animals were analysed in total) for H3K27me3, H3K4me3, and CTCF at the α-globin locus both in WT and D29 primary erythroid cells. Shaded grey bar indicates HS-29. The dashed box highlights the region over the Rhbdf1 and Mpg genes, magnified in top panels for ease of data visualisation.

Supplementary Figure 5 Effects of the individual deletion of HS-38 or HS-39 on local chromatin state in primary erythroid cells.

Normalised ATAC-seq and ChIP-seq read-densities (RPKM; 2 independent experiments in which 2 animals were analysed in total) for H3K27me3, H3K4me3, and CTCF at the α-globin locus in WT, D38 and D39 primary erythroid cells. Shaded grey bar indicates HS-38 and HS-39 (mutated CTCF sites in D38 and D39). The dashed box highlights the region over the Rhbdf1 and Mpg genes, magnified in top panels for ease of data visualisation.

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Hanssen, L., Kassouf, M., Oudelaar, A. et al. Tissue-specific CTCF–cohesin-mediated chromatin architecture delimits enhancer interactions and function in vivo. Nat Cell Biol 19, 952–961 (2017). https://doi.org/10.1038/ncb3573

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