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. 2020 Apr 10;23(5):101046. doi: 10.1016/j.isci.2020.101046

Demarcation of Topologically Associating Domains Is Uncoupled from Enriched CTCF Binding in Developing Zebrafish

Yuvia A Pérez-Rico 1,2,3,4, Emmanuel Barillot 2, Alena Shkumatava 1,5,∗
PMCID: PMC7182764  PMID: 32334414

Summary

CCCTC-binding factor (CTCF) is a conserved architectural protein that plays crucial roles in gene regulation and three-dimensional (3D) chromatin organization. To better understand mechanisms and evolution of vertebrate genome organization, we analyzed genome occupancy of CTCF in zebrafish utilizing an endogenously epitope-tagged CTCF knock-in allele. Zebrafish CTCF shares similar facets with its mammalian counterparts, including binding to enhancers, active promoters and repeat elements, and bipartite sequence motifs of its binding sites. However, we found that in vivo CTCF binding is not enriched at boundaries of topologically associating domains (TADs) in developing zebrafish, whereas TAD demarcation by chromatin marks did not differ from mammals. Our data suggest that general mechanisms underlying 3D chromatin organization, and in particular the involvement of CTCF in this process, differ between distant vertebrate species.

Subject Areas: Biological Sciences, Chromosome Organization, Molecular Biology

Graphical Abstract

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Highlights

  • •

    Identification of CTCF occupancy in zebrafish embryos using a tagged ctcf allele

  • •

    CTCF binding at promoters correlates with gene expression levels

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    No general CTCF enrichment at topological domain boundaries in zebrafish embryos

Biological Sciences; Chromosome Organization; Molecular Biology

Introduction

CCCTC-binding factor (CTCF) is a key regulator of gene expression and plays a central role in 3D organization of mammalian genomes (Dixon et al., 2012, Guo et al., 2015, Nora et al., 2017). In mammals, CTCF demarcates topologically associating domain (TAD) boundaries, and disruption of the CTCF sites in these regions results in the formation of ectopic contacts between neighboring domains (Despang et al., 2019, Dowen et al., 2014, Lupiáñez et al., 2015). CTCF and its role in gene regulation are conserved throughout bilaterians (Heger et al., 2012), whereas CTCF functions in 3D genome organization have diverged between invertebrates and vertebrates. In contrast to mammals, Drosophila CTCF is not essential for embryogenesis and its binding is not enriched at TAD boundaries (Gambetta and Furlong, 2018, Rowley et al., 2017). Given the functional divergence of CTCF in Drosophila and mammals, CTCF analyses in other non-mammalian vertebrate species are key for understanding the evolution and regulation of the 3D chromatin organization. Although the functions of CTCF in zebrafish development have been previously explored (Carmona-Aldana et al., 2018, Delgado-Olguín et al., 2011, Meier et al., 2018, Rhodes et al., 2010), no genome-wide CTCF in vivo binding data have been achieved in zebrafish so far. Similar to mammals, predicted CTCF binding motifs are distributed in divergent orientation at TAD boundaries in zebrafish (Gómez-Marín et al., 2015, Kaaij et al., 2018), suggesting the conserved role of CTCF in TAD demarcation. Here, we identified and characterize CTCF occupancy in developing zebrafish embryos using an epitope-tagged allele of ctcf. Although several gene regulatory features of zebrafish CTCF are similar to mammals, no enrichment of the in vivo CTCF occupancy was detected at TAD boundaries in zebrafish embryos, suggesting functional differences of CTCF in 3D genome architecture between vertebrates.

Results and Discussion

Identification of In Vivo CTCF Binding Sites Using the ctcfHPSH Zebrafish Allele

To determine CTCF occupancy in the zebrafish genome, we generated a tagged allele of ctcf, where a tripartite HA-PreScission-His tag was inserted in frame after the start codon of ctcf resulting in N-terminally endogenous CTCF tagged by HPSH (ctcf HPSH allele) (Figures 1A and S1A and Transparent Methods). We confirmed the expression of the tagged protein in ctcf HPSH/HPSH zebrafish (Figure 1B). Homozygous ctcf HPSH/HPSH zebrafish developed normally and were viable and fertile, indicating that the function of CTCF was not affected by the tag (Figures S1B and S1C). Chromatin immunoprecitation sequencing (ChIP-seq) analyses of CTCF binding in 24 hours postfertilization (hpf) ctcf HPSH embryos showed high correlation between biological replicates (Figures 1C and S1D) and confirmed a previously reported autoregulatory binding of CTCF to its promoter (Figure S1E) (Pugacheva et al., 2006). In the ChIP-seq data merged from both replicates, we identified 36,540 CTCF peaks that showed higher phastCons sequence conservation than random control regions (Figure 1D and Transparent Methods); the same trend was observed when considering only CTCF peaks that do not overlap exons (Figure S1F). Notably, the number of CTCF peaks identified in the zebrafish genome roughly corresponds to the number of CTCF sites in mammalian genomes (Pugacheva et al., 2020). Therefore, the ctcf HPSH zebrafish allele enables reliable and reproducible detection of the in vivo CTCF occupancy in the zebrafish genome.

Figure 1.

Figure 1

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Identification of CTCF Binding in Zebrafish

(A) Schematic representation of the ctcfHPSH zebrafish allele. Orange and purple boxes represent the inserted sequence and exons, respectively.

(B) Western blot using anti-hemagglutinin (HA) antibody on extracts from wild-type (WT) and ctcfHPSH/HPSH whole embryos and adult brains. Molecular weights are indicated on the right. γ-Tubulin served as a loading control.

(C) Tracks showing examples of CTCF peaks (purple bars) at the neurod2 and mycla loci (both located on the reverse strand). Displayed signal distributions and peaks correspond to biological replicates (Rep 1, Rep 2). Signal is represented on the y axis as –log10 (p value) of the CTCF ChIP-seq signal.

(D) Distribution of the average sequence conservation of CTCF peaks and control regions using peak centers as reference point.

See also Figure S1 and Tables S1 and S2.

Common Features of CTCF Binding Sites in Vertebrates

Because of the function of mammalian CTCF in establishing enhancer-promoter interactions (Sanyal et al., 2012), we analyzed zebrafish CTCF binding with respect to histone modifications and DNA accessibility in 24-hpf zebrafish embryos (Aday et al., 2011, Bogdanović et al., 2012, Gehrke et al., 2015, Irimia et al., 2012, Ulitsky et al., 2011) (Tables S1 and S2 and Transparent Methods). We found that zebrafish CTCF peaks were enriched for poised (H3K4me1), active (H3K4me3, H3K27ac), and accessible chromatin (ATAC-seq), but not for inactive chromatin (H3K27me3) (Figure 2A). Furthermore, de novo motif discovery identified a 20-bp core motif that was present in 78% of CTCF peaks and showed more similarity to the human CTCF motif than to the human CTCFL or Drosophila CTCF motifs (Figure 2B and Data S1). As reported for other vertebrate species (Boyle et al., 2011, Filippova et al., 1996, Kadota et al., 2017, Rhee and Pugh, 2011, Schmidt et al., 2012), we also identified enriched CTCF upstream motifs, separated from the core motif by 8- or 12-bp spacers (Figures 2C and S2A). Similar to mammalian CTCF binding sites, a fraction of which propagated in the genome through retrotransposition of repeat elements (Schmidt et al., 2012), non-autonomous DNA transposons were significantly enriched on CTCF binding sites (Figures 2D, S2B, S2C, and Table S3). Taken together, our analyses show that zebrafish and mammalian CTCF binding sites share similar features.

Figure 2.

Figure 2

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Characterization of CTCF Peaks and Binding Sites

(A) Heatmap profiles of four histone marks and ATAC-seq data at CTCF peaks ranked by decreasing CTCF ChIP-seq signal over the displayed region. Normalized signal is shown in FPM (fragments per million mapped fragments) for ATAC-seq and in RPKM (reads per kilobase million) for histone marks. All datasets correspond to the 24-hpf zebrafish embryonic stage.

(B) Dendrogram representing hierarchical clustering results of CTCF and CTCFL motifs. In total, 28,538 of 36,540 CTCF ChIP-seq peaks contained at least one matching site to the zebrafish motif, compared with 7,360 of 36,540 control sequences. Information content of each position on the x axis is expressed in bits on the y axis. Ncor, normalized Pearson correlation.

(C) Histogram showing the number of co-occurrences of the CTCF core and upstream motifs at different spacing distances (6–25 bp). Non-significant enriched spacing distances are shown in gray, enrichments are shown in pink, and the highest enrichments are shown in red. Bottom, inferred upstream motifs using sequences matching to the reference motif at the indicated distances from the CTCF core motif. Information content of each position on the x axis is expressed in bits on the y axis.

(D) Top five DNA transposon types enriched for CTCF binding sites. The fraction of repeats overlapping at least one CTCF motif is shown on the y axis. For control regions, the mean and the standard deviation (error bars) calculated by bootstrap analyses are shown.

See also Figure S2, Tables S1–S3 and Data S1.

CTCF Abundance at Promoters Correlates with the Gene Expression Levels

Although CTCF peaks were mainly located in intronic and intergenic regions, ∼6% of the peaks were found within promoters (Figure 3A). In human, a fraction of CTCF sites in promoter and intragenic regions is engaged in loops that impact exon inclusion (Ruiz-Velasco et al., 2017), raising the possibility of an equivalent mechanism in zebrafish. Because the resolution of the available zebrafish high-throughput chromosome conformation capture (Hi-C) data (∼20 kb) (Kaaij et al., 2018) does not allow investigation of this type of looping interactions, we sought to determine distinctive features of genes with CTCF-bound promoters. All genes that showed CTCF binding at their promoters were classified in three categories based on the signal strength of CTCF peaks ranging from high (top 10% percentile) to low (bottom 10% percentile) (Figure 3B and Transparent Methods). We found a positive correlation between the presence of CTCF motifs and CTCF occupancy regardless of the site orientation relative to transcription (Figure S3A) (χ2 tests of independence, p value ≤ 7.9 × 10−10). The increased CTCF occupancy at promoter also positively correlated with increased gene expression (White et al., 2017) (Figure 3C) and DNA accessibility (Figure 3D). Notably, CTCF binding at promoters is not a mere reflection of permissive chromatin, as we also identified promoters with high ATAC-seq signal but no CTCF binding (Figure S3B). Genes with high CTCF promoter occupancy had high signals for histone marks associated with enhancers (H3K4me1, H3K27ac), active promoters (H3K4me3, H3K27ac), and transcriptional elongation (Figures S3C–S3F). By contrast, no correlation between repressive chromatin (H3K27me3) and CTCF abundance was found (Figure S3G), whereas enrichment of the H3K27me3 repressive mark was overall higher at CTCF-bound promoters than at promoters without CTCF peaks (Figure S3H). In summary, our findings suggest that CTCF binding at promoters generally correlates with chromatin states that favor transcription. This observation could be explained by CTCF playing a role in the generation of nucleosome-depleted regions (Nora et al., 2017) or reflect CTCF binding in specific cell types to prevent ectopic gene expression, as previously reported for cis-regulatory elements of runx1 in zebrafish (Marsman et al., 2014).

Figure 3.

Figure 3

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High Abundance of CTCF Binding at Promoters Associates with High Gene Expression Levels

(A) Distribution of CTCF peaks across different zebrafish genomic regions. Percentages represent the number of CTCF peaks for each category.

(B) Average CTCF ChIP-seq signal profiles over promoters. Each line represents one of the three gene categories defined by CTCF abundance at promoters (low, medium, high) or promoters with no CTCF peaks (no peak).

(C) Expression of the stratified gene categories and genes without CTCF peaks at promoters. Differences in distribution are denoted as significant (∗) and non-significant (n.s.) according to two-sided Wilcoxon rank-sum test (p value ≤ 1.5 × 10−5).

(D) Average ATAC-seq signal profiles over promoters of gene categories defined by CTCF abundance as explained in (B).

See also Figure S3.

No Enrichment of CTCF Binding at TAD Boundaries in Zebrafish Embryos

Next, we sought to investigate the role of CTCF in zebrafish 3D genome organization by analyzing its distribution at TAD boundaries. Visualization of the available 24-hpf Hi-C data (Kaaij et al., 2018) showed enriched interactions of centromeres and telomeres and an uneven distribution of the signal along chromosomes (Figure S4A), which was even more pronounced at earlier developmental stages (Figure S4B). Although this signal distribution reflects the Rabl organization of chromosomes with continuous arm pairing over the cell cycle characteristic for dividing cells (Stadler et al., 2017) and the cell cycle heterogeneity of 24-hpf embryos (Figures S4B and S4C), we, nevertheless, identified 1,307 TADs (median size = 580 kb) using insulation scores (Crane et al., 2015) (Figure S4D and Table S4 and Transparent Methods). The latter was possible given that the 24-hpf Hi-C maps are a composite of the interactions occurring in dividing and interphase cells, which are characterized by lack and presence of TADs, respectively.

Although the annotated CTCF motif was enriched within accessible chromatin sites at TAD boundaries, this enrichment was not boundary specific, as it was also found enriched at accessible sites within TADs (enrichment p values < 1 × 10-20) (Figure S5A). Our analysis showed a moderate enrichment of in silico predicted CTCF sites and their divergent orientation bias within accessible chromatin at TAD boundaries, consistent with previous reports (Gómez-Marín et al., 2015, Kaaij et al., 2018) (Figures 4A and S5B). Similar to mouse CTCF (Dixon et al., 2012), only a small fraction of zebrafish CTCF peaks was located at TAD boundaries (Figure S5C). However, unlike enriched CTCF binding at TAD boundaries in mammals (Figure 4B), neither CTCF peaks nor the in vivo identified CTCF motifs were enriched at TAD boundaries in 24-hpf zebrafish embryos (Figures 4C and S5D). To exclude analysis bias, we applied the reciprocal insulation method and identified hierarchical domains (Zhan et al., 2017). In contrast to mammalian CTCF and in agreement with our zebrafish above-mentioned results, no reciprocal insulation value at which zebrafish CTCF enrichment was clearly maximized was identified (Figures S5E and S5F and Transparent Methods), potentially reflecting low variability in sequence composition of the genome (Costantini et al., 2007). Likewise, we found no zebrafish CTCF enrichment at boundaries of domains identified at 72.5% reciprocal insulation (i.e., the value at which the highest percentage of boundaries overlaps with CTCF peak summits) (Figure S5G). In agreement with the previous report (Kaaij et al., 2018), we found that TAD boundaries in zebrafish were enriched for chromatin marks associated with active transcription (Figures 4D and 4E), depleted for the H3K27me3 repressive mark (Figure 4F), and showed no enrichment for H3K27ac (Figure S5H) suggesting that the biochemical features of zebrafish TAD boundaries are similar to mammals. Our results do not imply that CTCF is dispensable for TAD establishment in the zebrafish genome, as we found moderate enrichment of predicted CTCF motifs within accessible chromatin at TAD boundaries and motif orientation biases, but they rather indicate that, in contrast to mammals, there is no strong correlation between TAD boundaries and high enrichment of CTCF. Importantly, it will require further investigations to determine if this moderate enrichment of CTCF is sufficient to establish TAD boundaries in zebrafish. It is also reasonable to propose that additional architectural proteins or the active chromatin state may play a role in TAD establishment in zebrafish, similar to Drosophila and other eukaryotes lacking this architectural protein. Interestingly, replication timing that is tightly associated with TAD distribution correlates with transcriptional status in zebrafish (Siefert et al., 2017) supporting the latter hypothesis. Moreover, it will be important to investigate colocalization of CTCF and cohesin binding in zebrafish, as the N-terminal CTCF region mediating the interaction with cohesin in mammals differs in its amino acid composition in zebrafish (Li et al., 2020, Pugacheva et al., 2006, Pugacheva et al., 2020). Indeed, this N-terminal region is highly conserved in organisms, in which CTCF is enriched at TAD boundaries including mammals and chicken (Fishman et al., 2018), but it is not conserved in Drosophila, in which CTCF does not delineate TAD boundaries (Moon et al., 2005, Rowley et al., 2017). Therefore, possible differences in the interaction between CTCF and cohesin in zebrafish may explain lack of CTCF enrichment at TAD boundaries. CTCF/cohesin ChIP-seq and Hi-C analyses in specific cell types and using single-cell approaches will be required to further investigate the functions of CTCF in the higher-order organization of the zebrafish genome.

Figure 4.

Figure 4

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Active Chromatin Marks but Not CTCF Are Enriched at TAD Boundaries

(A) Distribution of predicted CTCF motifs within ATAC-seq peaks (purple) along 600-kb regions centered on TAD boundaries (x axis) in 24-hpf zebrafish. The y axis shows the percentage of total peak counts in the 600-kb region located at each genomic position. The dashed line represents the mean background distribution, and the gray ribbon depicts the ±1 standard deviation range from the mean. Differences in mean percentages (central 60 kb) were assessed by Z scores. Non-significant, n.s.; ∗p < 1 × 10−5; ∗∗p < 1 × 10−20.

(B) Distribution of CTCF peaks (orange) relative to TAD boundaries identified in mouse embryonic stem cells as described in (A).

(C) Distribution of CTCF peaks (orange) relative to TAD boundaries identified in 24-hpf zebrafish embryos as described in (A).

(D) Distribution of H3K4me3-enriched peaks along TAD boundaries as described in (A).

(E) Distribution of H3K36me3-enriched peaks along TAD boundaries as described in (A).

(F) Distribution of H3K27me3-enriched peaks along TAD boundaries as described in (A).

See also Figures S4, S5, and Table S4.

Limitations of the Study

Although we found a positive correlation between CTCF binding at promoters and elevated gene expression, the cellular heterogeneity of 24-hpf zebrafish embryos does not allow to distinguish between CTCF-facilitating gene expression in specific cell types while acting as an insulator in other cells. Future cell-type-specific depletion of CTCF followed by purification of these cells will be required to interrogate CTCF binding and gene expression changes. In addition, it will also be important to analyze the relationship between CTCF enrichment and TAD boundaries in specific cell types to establish whether lack of strong CTCF enrichment at boundaries is maintained across different cell types or is cell specific.

Methods

All methods can be found in the accompanying Transparent Methods supplemental file.

Resource Availability

Lead Contact

Further information and requests for resources should be directed to and will be fulfilled by the Lead Contact, Alena Shkumatava (alena.shkumatava@curie.fr).

Materials Availability

The ctcf HPSH/HPSH zebrafish line generated in this study is available from the Lead Contact without restriction.

Data and Code Availability

CTCF ChIP-seq sequencing data generated in this study are available in the NCBI Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/). The accession number for the sequencing data reported in this study is NCBI GEO: GSE133437. No previously unreported algorithms were used to generate the results.

Acknowledgments

We thank members of the Shkumatava laboratory for discussions. ChIP-seq libraries generated in this study were sequenced at the ICGex NGS platform of Institut Curie funded by grants from “L’Agence Nationale de la Recherche”, France (ANR-10-EQPX-03 (Equipex) and ANR-10-INBS-09-08 (France Génomique Consortium) from the “Investissements d’Avenir” program) and by the Canceropole Île-de-France, France. This research was funded by grants from the European Research Council (FLAME-337440), “Fondation pour la Recherche Médicale”, France (DBI201312285578), and LabEx DEEP, France (ANR-11-LABX-0044, ANR-10-IDEX-0001-02).

Author Contributions

Conceptualization, Y.A.P.-R.; Methodology, Y.A.P.-R.; Formal Analysis, Y.A.P.-R.; Investigation, Y.A.P.-R.; Resources, E.B. and A.S.; Data Curation, Y.A.P.-R.; Writing – Original Draft, Y.A.P.-R.; Writing, Review & Editing, Y.A.P.-R and A.S.; Visualization, Y.A.P.-R.; Supervision, E.B. and A.S.; Funding Acquisition, E.B. and A.S.

Declaration of Interests

The authors declare no competing interests.

Published: May 22, 2020

Footnotes

Supplemental Information can be found online at https://doi.org/10.1016/j.isci.2020.101046.

Supplemental Information

Document S1. Transparent Methods, Figures S1–S5, Data S1, and Tables S1–S3
mmc1.pdf (1.4MB, pdf)
Table S4. Genomic Coordinates of Regions Used to Filter Topologically Associating Domains, Related to Figure 4
mmc2.xlsx (12.5KB, xlsx)

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Document S1. Transparent Methods, Figures S1–S5, Data S1, and Tables S1–S3
mmc1.pdf (1.4MB, pdf)
Table S4. Genomic Coordinates of Regions Used to Filter Topologically Associating Domains, Related to Figure 4
mmc2.xlsx (12.5KB, xlsx)

Data Availability Statement

CTCF ChIP-seq sequencing data generated in this study are available in the NCBI Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/). The accession number for the sequencing data reported in this study is NCBI GEO: GSE133437. No previously unreported algorithms were used to generate the results.

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