Abstract
Topologically Associating Domains (TADs) have been proposed to both guide and constrain enhancer activity. Shh is located within a TAD known to contain all its enhancers. To investigate the importance of chromatin conformation and TAD integrity on developmental gene regulation, we have manipulated the Shh TAD – creating internal deletions, deleting CTCF sites including those at TAD boundaries, as well as larger deletions and inversions of TAD boundaries. Chromosome conformation capture and fluorescence in situ hybridisation assays were used the investigate changes in chromatin conformation that result from these manipulations. Our data suggest that the substantial alteration of TAD structure has no readily detectable effect on Shh expression patterns during development – except where enhancers are deleted - and results in no detectable phenotypes. Only in the case of a larger deletion of one TAD boundary could some ectopic influence of the Shh limb enhancer be detected on a gene - Mnx1 in the neighbouring TAD. Our data suggests that, contrary to expectations, the developmental regulation of Shh expression is remarkably robust to TAD perturbations.
Introduction
At the megabase-scale, the mammalian genome is partitioned into self-interacting topologically associated domains (TADs) (Dixon et al., 2012; Nora et al., 2012) Mammalian TAD boundaries are enriched in CTCF sites in a convergent orientation (Narendra et al., 2015; Rao et al., 2014; Sanborn et al., 2015). TADs are formed by dynamic cohesin-driven loop extrusion (Fudenberg et al., 2016; Nora et al., 2017; Rao et al., 2017; Schwarzer et al., 2017; Vian et al., 2018) and convergent CTCF sites act to impede loop extrusion allowing WAPL to release cohesin from the chromosome (Haarhuis et al., 2017).
The regulatory landscapes of developmental genes are frequently found to be contained together in the same TAD (Dixon et al., 2012; Rao et al., 2014). TADs have therefore been proposed to act as functional regulatory units that favour contacts between enhancers and their target gene within a TAD whilst limiting aberrant interactions of enhancers across TAD boundaries (Fudenberg et al., 2016; Sun et al., 2019). In support of this, some studies have found that deletion or inversion of CTCF sites at TAD boundaries, can promote TAD boundary crosstalk and re-wire enhancer-promoter contacts (de Wit et al., 2015; Guo et al., 2015; Narendra et al., 2015; Rodríguez-Carballo et al., 2017). Moreover, a number of recent studies have suggested that changes to TAD structure can disrupt gene regulation through enhancer-rewiring in human disease (Flavahan et al., 2016; Franke et al., 2016; Lupiáñez et al., 2015).
However, other studies report that, although depletion of CTCF erases the insulation between TADs, it has limited effects on gene expression (Nora et al., 2017; Soshnikova et al., 2010). To further study the CTCF mediated function of TADs in developmental gene regulation, we have exploited the sonic hedgehog (Shh) regulatory domain – a paradigm locus for long-range regulation. The SHH morphogen controls the growth and patterning of many tissues during embryonic development, including the brain, neural tube and the limb. Spatial and temporal Shh expression is regulated by tissue-specific enhancers located within the gene, and in a large gene desert upstream of the gene (Anderson and Hill, 2014). Shh and its cis-acting elements are all contained within a well-characterised ∼960kb TAD (Anderson et al., 2014; Williamson et al., 2016). In the developing limb bud, Shh expression is solely determined by the ZRS enhancer (Sagai, 2005) located within an intron of the widely expressed Lmbr1, located 850kb upstream of Shh. Fluorescence in situ hybridisation (FISH) has shown that Shh and the ZRS are consistently located in relatively close proximity to each other in all cell types and tissues examined which we infer to be a consequence of the underlying invariant TAD structure. In contrast, we have observed increased ZRS-Shh colocalisation only in the Shh-expressing posterior portion of developing limb buds (Williamson et al., 2016). This might be consistent with a specific gene-enhancer contact.
To investigate the importance of chromatin architecture on TAD structure and thus on the regulation of gene expression, here, we extensively manipulate the Shh TAD and its TAD boundaries. We use a chromosome conformation assay (5C) and FISH to investigate how these manipulations affect TAD structures and interactions within and between TADs and we determine how these alterations affect the expression pattern of Shh and other developmentally regulated genes nearby. We also examine the phenotypic consequences of these manipulations. Our results question how important TADs are for correct spatial and temporal gene regulation.
Materials and Methods
Cell Culture and CrispR/cas9 mediated deletions
E14TG2A mouse embryonic stem cells (ESCs) were cultured under standard conditions (Anderson et al., 2014). CrispR guides were made by cloning annealed oligos (Table S1) into px458 (Addgene). 2μg of vector DNA were transfected into 8×105 ESCs using Lipofectamine 2000 (ThermoFisher) following the manufacturer’s instructions. After 48 hours, GFP positive cells were sorted by FACS and plated at low density. Ten days later, individual clones were picked and screened for correct deletion by PCR and Sanger sequencing (primers are listed in supplementary material Table S1).
Mouse lines and embryo analysis
The ShhΔ700 deletion was created by crossing the line SBLac96 (Anderson et al., 2014) to a line carrying a pCAGGS-Cre recombinase gene (Araki et al., 2006). With the exception of the Δ35kb and Inv35kb mouse lines, which were made by injection of the ESCs in to blastocysts, all of the other mouse lines were created as in Lettice et al., (2017) by direct microinjection into C57Bl6/ CBA F2 zygotes of the same guides as were used in ESCs. Resultant G0 mice are screened by PCR using flanking primers (Supp Table 1) and the deletions confirmed by Sanger sequencing. Lines were then established by crossing founder mice to C57Bl6 wildtypes.
LacZ expression analysis, in situ hybridisations and RT-PCR reactions were conducted as in Anderson et al. (2014).
FISH
E11.5 embryos were collected, fixed, embedded, sectioned, antibody stained for SHH expression and processed for FISH as previously described (Morey et al., 2007, Lettice et al., 2014), except that sections were cut at 8 μm. Fosmid clones (Figure 1A, Table S3) were prepared and labelled as previously described (Morey et al., 2007). Between 160-240 ng of biotin- and digoxigenin-labelled fosmid probes were used per slide, with 16-24 μg of mouse Cot1 DNA (Invitrogen) and 10 μg salmon sperm DNA. For 4-colour FISH, similar quantities of the additional fosmid was labelled with either Green496-dUTP (Enzo Life Sciences) or red-dUTP (Alexa FluorTM 594-5-dUTP, Invitrogen).
For 3D FISH on ESCs, 1×106 cells were seeded on slides for overnight. Cells were fixed in 4% paraformaldehyde (pFA) for 10 mins at room temperature and then permeabilized using 0.5% TritonX for 10 mins (Eskeland et al., 2010).
Image analysis
Slides were imaged using a Photometrics Coolsnap HQ2 CCD camera and a Zeiss AxioImager A1 fluorescence microscope with a Plan Apochromat 100x 1.4NA objective, a Nikon Intensilight Mercury based light source (Nikon UK Ltd, Kingston-on-Thames, UK) and either Chroma #89014ET (3 colour) or #89000ET (4 colour) single excitation and emission filters (Chroma Technology Corp., Rockingham, VT) with the excitation and emission filters installed in Prior motorised filter wheels. A piezoelectrically driven objective mount (PIFOC model P-721, Physik Instrumente GmbH & Co, Karlsruhe) was used to control movement in the z dimension. Step size for z stacks was set at 0.2 μm. Hardware control, image capture and analysis were performed using Nikon Nis-Elements software (Nikon UK Ltd, Kingston-on-Thames, UK). Images were deconvolved using a calculated point spread function with the constrained iterative algorithm of Volocity (Perkinelmer Inc, Waltham, MA). Image analysis was carried out using the Quantitation module of Volocity (Perkinelmer Inc, Waltham, MA).
3C library preparation
Limbs buds and bodies (with the limbs and heads removed) from wild type embryos, and entire ShhΔ700/Δ700 embryos were dissected at E11.5 and the tissue dissociated by pipetting in just enough PBS to cover them. The cells were fixed with 1% formaldehyde for 10 min at room temperature. For ESCs, 5×106 – 1×107 cells were fixed. Crosslinking was stopped with 125 mM glycine, for 5 min at r.t. followed by 15 min on ice. Cells were centrifuged at 400 g for 10 min at 4°C, supernatants removed, and cell pellets flash frozen on dry ice before storage at −80°C.
Cell pellets were treated as previously described (Dostie and Dekker, 2007; Ferraiuolo et al., 2010; Williamson et al., 2014). HindIII-HF (NEB) was the restriction enzyme used to digest the crosslinked DNA.
5C primer and library design
5C primers covering the Usp22 (mm9, chr11: 60,917,307-61,003,268) and Shh regions (mm9, chr5: 28,317,087-30,005,000) were designed using ‘my5C.primer’ (Lajoie et al., 2009) with the following parameters: optimal primer length of 30 nt, optimal TM of 65°C, default primer quality parameters (mer:800, U-blast:3, S-blasr:50). Primers were not designed for large (>20 kb) and small (<100 bp) restriction fragments, for low complexity and repetitive sequences, or where there were sequence matches to >1 genomic target. The Usp22 region was used to assess the success of each 5C experiment but was not used for further data normalization or quantification.
The universal A-key (CCATCTCATCCCTGCGTGTCTCCGACTCAG-(5C-specific)) and the P1-key tails ((5C-specific)-ATCACCGACTGCCCATAGAGAGG) were added to the Forward and Reverse 5C primers, respectively. Reverse 5C primers were phosphorylated at their 5′ ends. An alternating design consisting of 365 primers in the Shh region (182 Forward and 183 Reverse primers) was used. Primer sequences are listed in Table S9.
5C library preparation
5C libraries were prepared and amplified with the A-key and P1-key primers as described in (Fraser et al., 2012). Briefly, 3C libraries were first titrated by PCR for quality control (single band, absence of primer dimers, etc.), and to verify that contacts were amplified at frequencies similar to that usually obtained from comparable libraries (same DNA amount from the same species and karyotype) (Dostie and Dekker, 2007; Dostie et al., 2007; Fraser et al., 2010). We used 1 - 10 μg of 3C library per 5C ligation reaction.
5C primer stocks (20 μM) were diluted individually in water on ice and mixed to a final concentration of 2 nM. Mixed diluted primers (1.7 μl) were combined with 1 μl of annealing buffer (10X NEBuffer 4, New England Biolabs Inc.) on ice in reaction tubes. 1.5 μg salmon testis DNA was added to each tube, followed by the 3C libraries and water to a final volume of 10 μl. Samples were denatured at 95°C for 5 min and annealed at 55°C (48°C ESCs) for 16 hours. Ligation with Taq DNA ligase (10 U) was performed at 55°C (48°C ESCs) for one hour. One tenth (3 μl) of each ligation was then PCR-amplified individually with primers against the A-key and P1-key primer tails. We used 26 cycles based on dilution series showing linear PCR amplification within that cycle range. The products from 3 to 5 PCR reactions were pooled before purifying the DNA on MinElute columns (Qiagen).
5C libraries were quantified by bioanalyser (Agilent) and diluted to 26 pmol (for Ion PGM™ Sequencing 200 Kit v2.0). One microlitre of diluted 5C library was used for sequencing with an Ion PGM™ Sequencer. Samples were sequenced onto Ion 316™ Chips following the Ion PGM™ Sequencing 200 Kit v2.0 protocols as recommended by the manufacturer (Life TechnologiesTM).
5C data analysis
Analysis of the 5C sequencing data was performed as described in (Berlivet et al., 2013). The sequencing data was processed through a Torrent 5C data transformation pipeline on Galaxy (https://main.g2.bx.psu.edu/). Before normalizing, interactions between adjacent fragments were removed due to the high noise: signal ratio likely to occur here. Data was normalized by dividing the number of reads of each 5C contact by the total number of reads from the corresponding sequence run. All scales shown correspond to this ratio multiplied by 103. The number of total reads and of used reads is provided for each experiment in Table S10. 5C datasets are to be uploaded to the Gene Expression Omnibus (GEO) website http://www.ncbi.nlm.nih.gov/geo/.
RESULTS
A large deletion within the Shh TAD does not disrupt local genome organisation or limb-specific activation of Shh
Prominent architectural features of the Shh TAD include CTCF bound at five binding sites at both boundaries across multiple cell types (Fig. 1A), and two sub-TADs with overlapping boundaries located within the gene desert between the forebrain enhancers and Rnf32 (Figs. 1F and 2A heat maps). This region of the gene desert includes less well defined CTCF peaks that are not invariant across cell types but due to their location may have some role in defining these sub-TADs (Fig. 1A) (Rosenbloom et al., 2013).
To determine the contribution of TAD internal sequence to 3D chromatin organisation and gene expression, we exploited our previous work that used the local hopping activity of the sleeping beauty (SB) transposon to probe the Shh regulatory domain (Anderson et al., 2014). Transposition of the SB leaves a LoxP site at the initial integration site and inserts a second LoxP site where it re-integrates, enabling Cre recombinase to create deletions of the intervening DNA. The orientation of the re-integration means the LacZ gene carried by the SB is retained in the deleted chromosome allowing remaining enhancer activity to be monitered. Using this approach, we deleted approximately 700kb (∼70%) of the internal Shh TAD sequence, including the sub-TAD boundaries, but, leaving the five CTCF binding sites at the extremes of the TAD intact (Fig. 1A). The Δ700 deletion removes many of the known Shh enhancers, and relocates the ZRS to within 96kb of the Shh promoter (Fig. 1A). Removal of the Shh forebrain and epithelial enhancers in the Δ700 deletion is shown by LacZ staining of ShhΔ700/+ embryos which shows staining only within the floor plate and hind brain, presumably driven by the proximal enhancers SFPE1/2 and SBE1, and within the limbs driven by the ZRS (Figs. 1B & C). Homozygous ShhΔ700/Δ700 embryos show phenotypes very similar to those of Shh-/- embryos but with normal limb and digit patterning (Chiang et al., 1996). These data indicate that, despite its incorrect position now only 96kb from the Shh promoter, ZRS is able to function normally to drive Shh expression in limb development (Figs. 1D & 1E).
To determine how removal of this ∼700kb disrupts local TAD structure we carried out 5C on whole E11.5 Shh Δ700/ Δ700 and wild type embryos. 5C heatmaps show that the Shh TAD boundaries and the adjacent TADs are unaffected by the deletion, (Figs. 1F, 1G & S1). Together, these data show that interactions within the deleted region are not required for ZRS activity or for maintaining the location of the TAD boundaries, and there is not a requirement for a great genomic distance between Shh and its limb enhancer.
Interactions within the Shh TAD are delineated by CTCF sites either side of Shh and within Lmbr1
Our previous 5C analyses on cells dissected from whole limbs, bodies and heads of E11.5 embryos showed enriched interactions between the genomic region containing Shh, located at one TAD boundary, and a genomic region within Lmbr1 close to ZRS, located ∼70kb from the other TAD boundary (Williamson et al., 2016). That this enrichment can be identified throughout the E11.5 embryo, a stage when we have shown that high levels of Shh-ZRS colocalisation occur only in the posterior distal limb, excludes active Shh-ZRS colocalisation as the sole driver of this apparent chromatin loop (Williamson et al., 2016). To gain further insight into the nature of these interactions, we dissected E11.5 limb buds to compare cell populations with no ZRS activity (anterior 2/3 of bud) with those with ZRS-active cells (posterior 1/3) (Figs. 2A and S2A).
The Shh TAD structure revealed by 5C is similar in both anterior and posterior limb bud cell populations, and comparable to dissected E11.5 bodies (compare Fig. 1F and Fig. 2A). At high (15kb) resolution, the strongest enrichment in both populations involved interactions between both Shh and the genomic region immediately 3′ of Shh, and a locus ∼20kb from ZRS in intron 5 of Lmbr1 (Figs. 2B & S2B, left- and right-hand heatmaps). ENCODE data (Rosenbloom et al., 2013) indicates these three loci are all bound by CTCF across a range of cell and tissue types (Fig. 1A), with the underlying DNA containing CTCF-binding motifs in a convergent orientation suggestive of roles in blocking loop extrusion (Fig. 2A). There is a subtle enrichment of interactions apparently between Shh and ZRS in the posterior population which contains ZRS-active cells, known as the Zone of Polarising Activity (ZPA) (Figs. 2B & S2B, centre heatmap). The minimal difference between the anterior and posterior tissue could be due to the presence of cells in the posterior population where ZRS is not active diluting the interactions between Shh and ZRS.
Deletion of CTCF sites at Shh reduces Shh intra-TAD interactions and disrupts Shh/ZRS proximity
The CTCF-anchored loop located at convergent binding sites near to both Shh and ZRS detected by 5C throughout the E11.5 embryo would appear to be important for maintaining Shh/ZRS spatial proximity and possibly contributes to the formation of the Shh TAD along with the CTCF binding sites located at the Lmbr1 promoter (Fig. 1A). We therefore used CRISPR-Cas9 to delete sequences containing these domains of CTCF binding in mouse ESCs (The size for each individual deletion is listed in Table S1). We first deleted the CTCF binding regions 3′ and 5′ of Shh (sites numbered 1 and 2 respectively in Fig. 1A). We generated ESC lines homozygous for each of these deletions and compared chromatin conformation by 5C and FISH.
The Shh TAD structure in ESCs is similar to that in E11.5 embryos (Figs. 3A & S3A, top row, left-hand 5C heatmaps). Deletion of CTCF site 1 (ΔCTCF1), which delineates the TAD boundary 3′ of Shh, re-locates the TAD boundary by ∼40kb to 5′ of Shh to the vicinity of CTCF2 (Figs. 3A & S3A, top row, centre heatmaps). This is evidenced by Shh losing interactions with the rest of its own TAD in ΔCTCF1 cells, and gaining interactions with regions just 5′ of En2 and Rbm33 (Figs. 3A & S3A, bottom row, left-hand heatmaps). There is also loss of interactions with a locus upstream of the forebrain enhancers near the sub-TAD boundary within the larger Shh TAD. These data would be consistent with CTCF1 forming the Shh TAD boundary by blocking a loop extrusion process emanating from within the En2 TAD.
The left hand Shh TAD boundary is not affected by deletion of CTCF site 2 (Figs. 3A & S3A, top row, right-hand heatmaps), however, the 5′ Shh region does gain contacts with the En2 TAD in a similar manner to the loss of CTCF1 (Figs. 3A & S3A, bottom row, right-hand heatmaps), suggesting that both CTCF1 and CTCF2 are necessary to optimally block loop extrusion emanating from the En2 TAD. There are also strongly enriched interactions within the Shh sub-TADs detected in ΔCTCF2 cells (Figs. 3A & S3A).
We also analysed possible alterations of chromosome conformation due to the CTCF site deletions with 3D-FISH using probes for Shh, SBE2 and ZRS (Fig. 3B). Interprobe distances between all three probe pairs were significantly increased in the CTCF deletion cells compared to wild type ESCs (Fig. 3C), consistent with the reduced interactions between Shh and the rest of the TAD identified by 5C. Conversely, we detected significantly decreased distances between Shh and Cnpy1 (in the neighbouring En2 TAD) in ΔCTCF1 cells (but not ΔCTCF2) compared to wild type (Figure 3C), consistent with the relocation of the TAD boundary.
Deleting either CTCF1 or CTCF2 disrupts Shh-ZRS spatial proximity in ESCs and, more generally, result in reduced 5C interactions between Shh and the rest of the regulatory TAD that may be due to the re-location of the TAD boundary (ΔCTCF1) or greater sub-division of the TAD (ΔCTCF2). The TAD boundary adjacent to Shh is sharply defined by CTCF1 whereas the boundary location of the neighbouring En2 TAD cumulatively results from both CTCF 1 and 2, possibly by blocking loop extrusion emanating from this TAD. However, neither of these deletions on their own is sufficient to cause the merging of the two and neighbouring TADs.
Shh-ZRS proximity is disrupted by the deletion of ZRS/Lmbr1 CTCF sites
Both CTCF1 and CTCF2 have highly enriched interactions with the CTCF site ∼20 kb from ZRS in intron 5 of Lmbr1 (CTCF3) (Figure 1A, site 3). Therefore, we deleted both alleles of CTCF3 (ΔCTCF3) to determine the consequences for chromosome conformation.
Whilst whole TAD integrity was unaffected by ΔCTCF3 (Figs. 4A & S4A, top row, centre-left heatmaps), intra-TAD reorganisation occurred in a similar manner to the loss of CTCF2, with enriched interactions within the sub-TADs (Figs. 4A & S4A). FISH inter-probe distances between Shh, SBE2 and ZRS (Fig. 4B) all significantly increased in ΔCTCF3 cells compared to wild type (Fig. 4C). These data suggest that loss of any one of the three CTCF binding domains (1, 2 or 3) can disrupt the spatial proximity of Shh, SBE2 and ZRS (Fig. 4C, Figure 3C).
Finally, we generated ESC lines with deletions of CTCF binding sites at the Lmbr1 promoter (ΔCTCF4) and 5′ Lmbr1 (ΔCTCF5), both of which are located at the boundary between the Shh TAD and the adjacent TAD containing Mnx1 (Fig. 1A). Neither of these deletions affected Shh TAD integrity or boundary positions (Figures 4A & S4A, top row, centre-right and right-hand heatmaps). However, the CTCF3/ZRS genomic region had enriched interactions (red) across the Shh TAD in ΔCTCF4 ESCs, especially with the Shh locus itself (Figs. 4A & S4A, bottom row, middle heatmaps). The CTCF motif within CTCF5 is oriented towards the adjacent Mnx1-containing TAD and deletion of the site caused a loss of interactions (blue) between this boundary region and the Mnx1 TAD (Figs. 4A & S4A, bottom row, right-hand heatmaps).
FISH revealed significantly increased inter-probe distances between Shh and ZRS in ΔCTCF5 cells and between SBE2 and ZRS in ΔCTCF4 cells, which contrasts with the 5C data (Fig. 4C). There are also decreased distances seen between ZRS and Mnx1 in the adjacent TAD in the absence of CTCF4, something not apparent in the 5C data (Fig 4D). Also, in contrast to 5C data, loss of CTCF5 decreases distances between ZRS and Lmbr1 promoter compared to wild type (Fig. 3D).
We conclude that deletion of CTCF binding sites at either of the Shh TAD and sub-TAD boundaries, especially CTCF1, 2 and 3, affects local chromatin organisation in ESCs and disrupted Shh/ZRS spatial proximity.
Reduced Shh-ZRS colocalisation upon the loss of CTCF1, 2 and 3 in the limb
To test how disrupted TAD organisation impacts on chromosome conformation and Shh gene expression during embryonic development, we generated mouse lines carrying each of the CTCF deletions. We previously reported a significantly enhanced Shh-ZRS colocalisation in the Zone of Polarising Activity (ZPA) of the limb bud where Shh is active compared to non-expressing limb tissues. This colocalisation depends on a fully functional ZRS and, when mutations are made within ZRS that deleteriously affect Shh expression, colocalisation rates are dramatically reduced (Lettice et al., 2014; Williamson et al., 2016). Therefore, we assayed the spatial proximity of Shh, SBE2 and ZRS by FISH in E11.5 mouse embryo sections that include posterior (ZPA) and anterior distal limb tissue from wild-type and homozygous ΔCTCF mutant embryos (Fig. 5A).
In both regions of the wild type limb bud analysed (ZPA and anterior), Shh-ZRS distances were shorter, than between Shh-SBE2 and SBE2-ZRS, consistent with these two loci being maintained in spatial proximity across the limb bud (Figs. 5B & C, white violin plots). Distances between Shh and both SBE2 and ZRS were significantly increased in ΔCTCF1, 2 and 3, but not ΔCTCF4 and 5 embryos, similar to that observed in ESCs (Figs. 5B & C). This disruption of Shh-ZRS spatial proximity in the ZPA of ΔCTCF1, 2, and 3 mutant embryos significantly reduced the frequency of Shh-ZRS colocalisation (<200nm) down to levels seen in non-expressing parts of the wild-type limb bud (Fig. 5D).
Shh expression patterns and development are unaffected in CTCF site deletion mice
Our data indicate that deletion of individual CTCF sites affects TAD boundaries, intra- and inter-TAD interactions and enhancer-promoter co-localisation frequencies. These alterations in 3D chromosome conformation would be predicted to have an effect on gene expression. Surprisingly, however, we found that mice homozygous for any of the ΔCTCF deletions are viable, fertile and have no detectable phenotype. In situ hybridisation in homozygous mutant embryos showed a normal pattern of Shh expression, and at similar levels to wild type, (Fig. 6A and B). At E11.5 expression is detected only within the developing midline of the brain, the Zli and the medial ganglionic eminence in the head and staining is visible in the floor plate and notochord, the ZPA of the limb buds and umbilicus in the body. No ectopic expression is detected at the midbrain / hindbrain junction driven by neighbouring En2 or Cnpy1 enhancers (Fig 6A), Conversely, in embryos homozygous for ΔCTCF1 and ΔCTCF2 there is no evidence for ectopic En2 and Cnpy1 expression in any of the normal sites of Shh expression in the brain (Fig 6C and D).
Similarly, no ectopic Shh expression is detected in motor neurons driven by Mnx1 enhancers in the TAD beyond ZRS/Lmbr1 (Fig. 6A), and Mnx1 was not expressed ectopically in any of the normal sites of Shh expression in embryos carrying homozygous deletions of CTCF3, CTCF4 and CTCF5 (Fig 6E). These findings indicate that enhancer/promoter specificity is maintained in these deletion embryos and that there is no cross-talk across TAD boundaries resulting in ectopic expression driven by Shh enhancers, even in the absence of these CTCF sites.
Mice heterozygous for a Shh null allele express only about 60% wild type levels of Shh but develop normally, suggesting that there is a wide range of Shh expression levels capable of driving normal development. Indeed, in the limb Shh levels must fall to about 20% of wildtype before development becomes perturbed and digits are lost (Lettice et al., 2017). We therefore made compound heterozygotes carrying both the ΔCTCF1 and Shh null alleles to uncover subtle effects on Shh expression caused by the CTCF1 deletion. These ShhΔCTCF1/- mice develop normally and are viable and fertile, further suggesting that deletion of CTCF1 results in no deleterious changes in Shh expression.
A 35kb deletion that removes the Lmbr1 promoter and TAD boundary disrupts chromatin conformation with no deleterious phenotype
Deletion of CTCF1 3′ of Shh showed that this position was important for the TAD boundary location and ensuring Shh remained in physical proximity with its regulatory domain (Figs. 3 & S3), even though the loss of this site had no apparent phenotypic consequence (Fig. 6). Deleting either CTCF4 or CTCF5 at the other TAD boundary had no effect on the Shh boundary location and resulted in a minimal loss of proximity between ZRS and Shh (Figs. 4 & S4). Loss of TAD boundary regions can result in the merging of adjacent TADs and the ectopic activation of genes in one TAD by enhancers in the other merged TAD, with phenotypic consequences (Fabre et al., 2017; Lupiáñez et al., 2015). However, this involved the deletion of sizeable stretches of DNA across the boundaries in question, tens of kilobases rather than individual CTCF sites. In addition to CTCF binding sites, a number of features are found enriched at TAD boundaries including those associated with active promoters (Dixon et al., 2012). To determine if a more extensive deletion across the Lmbr1 boundary results in the merging of adjacent TADs, a homozygous 35kb deletion (Δ35) was generated which removed CTCF4 and CTCF5, in all covering about 13kb upstream of the Lmbr1 TSS, as well as TSS itself and first two exons of Lmbr1 (Fig. 1A). RT-PCR confirmed that this deletion eliminates transcription throughout the 5′ end of Lmbr1 in both isolated limb buds and the rest of the body (Fig 7A).
5C showed that Δ35 caused the relocation of the TAD boundary a further ∼40kb 5′ of the Lmbr1 promoter towards the promoter of Nom1 (Figs. 7B & S5A & B), rather than a merging of the adjacent TADs. There are enriched interactions of the region from CTCF3 to Nom1 across the Shh TAD in Δ35 cells (red in Fig. 7C) compared to wild type and the CTCF3/ZRS genomic region gains interactions into the adjacent TAD up to Mnx1 (Figs. 7C & S5B), whereas the region around Nom1 had reduced interactions with its own TAD. Virtual 4C plots derived from the 5C data show that ZRS gains contacts with the rest of the Shh TAD and into the adjacent TAD up to Mnx1 (Fig. S5C, top graphs), and the new boundary region gains contacts with the Shh TAD while losing interactions with the Mnx1 TAD (Fig. S5C, bottom graphs). 3D-FISH (Fig. 7D) showed that distances between ZRS and the other three labelled loci (Shh, SBE2 and Mnx1) were all significantly decreased in Δ35 (Fig. 7E), which corresponds to the 5C and virtual 4C data. The reduced spatial distance between ZRS and Mnx1 was not due to the linear genomic distance being reduced because of the 35kb deletion, as similar effects were seen in cells carrying an inversion of this DNA rather than a deletion (Fig. 7E).
Removal of the Shh TAD boundary at the Lmbr1 promoter relocates the boundary to the promoter of Nom1, and the ZRS has enhanced ability to contact sequences both within its own TAD and the Mnx1 TAD. Despite these differences, Δ35 homozygous mice were viable, fertile and had no apparent phenotype. The Shh expression pattern is also indistinguishable from wild type (Figs. 7F & G) - in particular midline expression is detected in the floor plate and notochord as one stripe down the body (Fig 7G, arrow head), with no evidence for expression as two more lateral stripes driven by Mnx1 motor neuron enhancers (Fig. 6E). Even in the sensitised background of the Shh null chromosome compound ShhΔ35/- mice are also phenotypically normal.
Interestingly however, given the decreased distances measured by FISH between ZRS and Mnx1, the limb expression of Mnx1 was increased in ShhΔ35/Δ35 embryos in comparison to wild-type embryos (Figs. 7H-J). This suggests that this deletion, that encompasses the TAD boundary, enhances the ability of the Mnx1 promoter to respond to the ZRS. However, no upregulation of Mnx1 expression is seen in the pharyngeal endoderm and developing lungs which would be driven by the enhancers neighbouring ZRS, MACS1 and MFCS4.
DISCUSSION
A systematic genetic approach to remove most of the Shh enhancers (excluding ZRS) (Δ700), to delete individual CTCF sites, and to delete or invert large regions encompassing a TAD boundary, has enabled us to use chromosome conformation capture and imaging to assay the resulting perturbations to chromosome organisation within the Shh regulatory TAD, and between this and neighbouring TADs. Despite this, we detected little or no perturbation of gene regulation during embryonic development and no detectable phenotype in animals that can be attributed to this altered chromosome conformation.
ZRS activity is not distance dependent and does not require factors located within the intervening gene desert
5C analysis confirmed that TAD boundaries were unaffected by removal of most of the internal region of the Shh TAD (Δ700) (Figs. 1 & S1), with Shh and its remaining enhancers still located within the same, but smaller, TAD. This large deletion did cause extensive disruption to the developing embryo, mainly, it can be assumed, due to the loss of several known forebrain and epithelial enhancers within the deleted region. However, even in embryos homozygous for the 700kb deletion, which relocates ZRS to less than 100kb distant from Shh, ZRS function is maintained, there is no detrimental effects on limb bud-specific Shh activation and normal development of the limbs occurs. Therefore, the large genomic distance from Shh is not intrinsic to the function of the ZRS. This is in contrast to the loss of interactions following similar perturbations between a limb-specific enhancer and Hoxd13 that resulted in loss of Hoxd13 activity (Fabre et al., 2017).
Loss of CTCF sites at the Shh TAD boundaries disrupts chromatin architecture, and impacts Shh/ZRS spatial proximity
We have previously shown that Shh and ZRS are in spatial proximity (∼300nm) in the early embryo in both expressing limb tissue and the non-expressing adjacent flank (Williamson et al., 2016). Here, using 5C on cells dissected from E11.5 anterior and posterior limb buds we show that this is driven by a looping interaction between the sites 3′ and 5′ of Shh (containing CTCF1 and CTCF2 sites) and a region within intron 5 of Lmbr1 about 20kb from ZRS (CTCF3) (Figs. 2 & S2). This loop is also present in ESCs, and spatial proximity of Shh and ZRS is lost upon the deletion of any one of the three CTCF sites in both ESCs and E11.5 limb bud tissue (Figs. 3, 4, 5, S3 & S4). Deleting CTCF sites at the Lmbr1 promoter TAD boundary (CTCF4 and CTCF5) had less effect on Shh/ZRS spatial proximity. Increased inter-probe distances between either Shh or ZRS and the forebrain enhancer SBE2 located at the centre of the TAD suggest that the loss of spatial proximity may be due to a general decompaction throughout the TAD.
Shh responds to its developmental enhancers regardless of TAD disruption
5C analysis in ESCs suggests that the disruption caused by the deletions removes Shh from its regulatory TAD (ΔCTCF1) or re-enforces contacts within sub-TAD domains such that the forebrain enhancers are sequestered in one and ZRS and the long-range epithelial enhancers in the other, with a loss of interactions between both sub-TADs and either sub-TAD with Shh (ΔCTCF2 and ΔCTCF3). Nevertheless, in all of these configurations, the pattern of Shh during embryonic development appears to be normal and the resulting mice have no detectable phenotype. This indicates that communication between Shh and its extensive set of developmental enhancers is remarkably robust to TAD perturbation.
Ectopic expression across disrupted TAD boundaries is not common
Loss of CTCF1 not only moves the TAD boundary ∼40kb to beyond the 5′ end of Shh but also enables greater interactions between Shh the adjacent TAD which contains other genes and their enhancers active during brain development, but in a pattern distinct from Shh. En2 is expressed at the mid-hindbrain boundary, a pattern at least partly dependent on an enhancer binding Pax2/5/8 (Li Song and Joyner, 2000). Similarly, Cnpy1 expression at the mid-hindbrain boundary is thought to be important for FGF signalling (Hirate and Okamoto, 2006). Despite increased chromatin interactions over the Shh TAD boundary in ΔCTCF1, there is no ectopic expression of Shh in the mid-hindbrain driven by the En2/Cnpy1 enhancers and, vice versa, there is no ectopic expression of En2/Cnpy1 at sites driven by Shh enhancers (Fig. 6).
The Lmbr1 boundary has been suggested to be less precise than the Shh boundary from a structural and regulatory point of view (Symmons et al., 2016). Indeed, deletion of CTCF4 or 5 had little effect on this Shh TAD boundary. However, ΔCTCF5 weakened the boundary of the neighbouring Mnx1 TAD and increased proximity between ZRS and Mnx1 was detected in ΔCTCF4. However, in neither case was there evidence for enhanced expression of Mnx1 – e.g. in limb buds driven by ZRS – beyond that detected in wild-type embryos. Interestingly, even in wildtype situations, Mnx1 has a weak expression domain concomitant with the limb bud ZPA, suggesting that this gene may be influenced by ZRS activity emanating from the adjacent TAD. Nor was there evidence of the Mnx1 motor-neuron enhancer (Zelenchuk and Brusés, 2011) driving expression of Shh in motor neurons of the developing neural tube in any of the mutant embryos.
A larger (35kb) deletion of this boundary removing CTCF4, CTCF5 and the promoter/first two exons of Lmbr1, enhanced ZRS 5C contacts across both the Shh TAD and into the neighbouring Mnx1 TAD (Figs. 7 & S5). Increased Mnx1 expression in the ZPA of embryos homozygous for the 35kb deletion suggests that the potentially increased contacts between Mnx1 and ZRS identified in ESCs could be enabling greater activation of this gene by the Shh limb enhancer.
Perturbations of the Shh TAD boundaries can negatively impact on gene-enhancer co-localisation but are insufficient to cause a deleterious phenotype
It is commonly thought that enhancer driven gene-activation required ‘contact’ or very close apposition of the enhancer and promoter. Inversions encompassing the Shh TAD boundaries that disrupted TAD integrity and significantly increased the genomic distance between Shh and ZRS result in severe limb malformations, suggesting that these rearrangements prevent ZRS from contacting/regulating the Shh promoter (Symmons et al., 2016). These data and our 5C and FISH analysis which shows that the Shh TAD forms a compact, discrete regulatory hub (Williamson et al., 2016) suggest that 3D organisation of the Shh TAD could allow distal enhancers to come into close proximity to selectively regulate Shh expression. However, in the functionally relevant cells of the limb bud ZPA, ZRS colocalisation (<200 nm) with Shh was reduced to levels of the non-expressing distal anterior levels in ΔCTCF1, ΔCTCF2 and ΔCTCF3 homozygous embryos without adversely affecting Shh expression (Fig. 6) and with no subsequent phenotypical effects. We have previously shown that reduced Shh-ZRS colocalisation as a consequence of deleting the 3′ end of ZRS (which was shown to be not required for activating proximal expression of a reporter gene in the ZPA) caused the loss of Shh forelimb bud expression and severe attenuation of hindlimb bud expression which resulted in a range of limb malformations (Lettice et al., 2014). Therefore, loss of colocalisation on its own does not have severe enough effects on levels of expression to result in limb malformations.
All embryos homozygous for one of the five CTCF binding domain deletions and even with the 35kb deletion of the Lmbr1 boundary developed normally and were able to reproduce. Moreover, sufficient expression of Shh was maintained in compound heterozygote embryos carrying either ΔCTCF1 or the 35kb deletion opposite a Shh null allele for these mice to have no phenotype. A contemporaneous study on the same genomic territory has largely re-capitulated these results – deletions of Lmbr1 CTCF sites and the gene promoter caused perturbations to local chromatin conformation but Shh expression, although reduced, was enough to drive normal limb development (Paliou et al., 2019). The Shh regulatory landscape is set up to ensure optimal activation of the gene and here we have shown this is robust to perturbations of TAD integrity and structure. Only large-scale disruptions incorporating boundaries appear to cause TADs to merge with resulting developmental defects (Lupiáñez et al., 2015).
Our data suggest that TADs are largely structural and play no overt role in regulating gene expression. We speculate that the largely unvarying organisation of TADs could have provided the necessary stable genomic environment for the accumulation of regulatory elements over evolutionary time rather than being essential for target gene activation.
Acknowledgements
We thank the staff of the IGMM advanced imaging resource and technical services for their assistance with imaging and sequencing. We would also like to thank the staff at the BRF/Evans Building for expert technical assistance. This work was supported by the Medical Research Council, UK.