A PRE loop at the dac locus acts as a topological chromatin structure that restricts and specifies enhancer–promoter communication

Denaud, Sandrine; Bardou, Marion; Papadopoulos, Giorgio-Lucio; Grob, Stefan; Di Stefano, Marco; Sabarís, Gonzalo; Nollmann, Marcelo; Schuettengruber, Bernd; Cavalli, Giacomo

doi:10.1038/s41594-024-01375-7

A PRE loop at the dac locus acts as a topological chromatin structure that restricts and specifies enhancer–promoter communication

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Published: 16 August 2024

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A PRE loop at the dac locus acts as a topological chromatin structure that restricts and specifies enhancer–promoter communication

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Abstract

Three-dimensional (3D) genome folding has a fundamental role in the regulation of developmental genes by facilitating or constraining chromatin interactions between cis-regulatory elements (CREs). Polycomb response elements (PREs) are a specific kind of CRE involved in the memory of transcriptional states in Drosophila melanogaster. PREs act as nucleation sites for Polycomb group (PcG) proteins, which deposit the repressive histone mark H3K27me3, leading to the formation of a class of topologically associating domain (TAD) called a Polycomb domain. PREs can establish looping contacts that stabilize the gene repression of key developmental genes during development. However, the mechanism by which PRE loops fine-tune gene expression is unknown. Using clustered regularly interspaced short palindromic repeats and Cas9 genome engineering, we specifically perturbed PRE contacts or enhancer function and used complementary approaches including 4C-seq, Hi-C and Hi-M to analyze how chromatin architecture perturbation affects gene expression. Our results suggest that the PRE loop at the dac gene locus acts as a constitutive 3D chromatin scaffold during Drosophila development that forms independently of gene expression states and has a versatile function; it restricts enhancer–promoter communication and contributes to enhancer specificity.

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Main

Eukaryotic genomes are highly organized within the three-dimensional (3D) nuclear space. The development of chromatin conformation capture (3C)-based methods and advanced microscopy approaches highlighted the importance of the 3D chromatin topology of cis-regulatory elements (CREs) in gene regulation¹. Regulatory interactions of CREs define, maintain and change the expression pattern of key developmental genes to ensure proper development. Loss of these control mechanisms is a frequent feature of cancer and disease^2,3.

In interphase nuclei, chromosomes are hierarchically organized into topologically associating domains (TADs)^4,5,6 that modulate gene regulation, although their exact function is debated. Extensive rearrangement of chromosomes and TADs in the Drosophila genome does not correlate with changes in gene expression⁷ and removal of proteins involved in TAD border formation have no dramatic effect on gene expression⁸. On the other hand, genomic rearrangements of TADs can cause gene misexpression and diseases^{9,10,11,12,13}. TADs facilitate local promoter–enhancer interactions and prevent inappropriate interactions between different TADs^4,9,14. Genome organization can shape transcription dynamics by two complementary mechanisms; ‘tethering elements’ (TEs) within TADs foster interactions between CREs, whereas insulators and/or TAD boundaries prevent inappropriate interactions between enhancers and promoters¹⁵.

In Drosophila, a specific class of TADs is characterized by the presence of the H3K27me3 Polycomb mark, which can cover several hundreds of kilobases, including key developmental regulatory genes^16,17. Within these Polycomb domains, epigenetic regulatory sequences named Polycomb response elements (PREs) act as nucleation sites for the recruitment of the Polycomb repressive complexes 2 and 1 (PRC2 and PRC1), which are responsible for the deposition and spreading of H3K27me3 and H2AK118ub, respectively, as reviewed in the literature¹⁸. Many Polycomb domains contain multiple PREs and genes, which tend to be coregulated or involved in related developmental pathways^16,17. PREs can participate in the maintenance of both active and repressed gene expression states throughout development, as reviewed in the literature¹⁹, and a subset of PREs can engage in specific chromatin contacts within Polycomb domains, forming chromatin loops (PRE loops)^20,21,22. PRC1 is likely to have an important role in PRE looping because PRC1 is critical for chromatin condensation of Polycomb domains and the establishment of their long-range interactions in mammals, as reviewed in the literature¹⁸. Looping interactions might be mediated by oligomerization of the sterile alpha motif domain of Polyhomeotic (PH), a subunit of PRC1, which is crucial for the condensation of individual Polycomb domains²³ and for mediating long-range Polycomb domain interactions^24,25.

Using the Polycomb domain associated with the leg patterning gene dachshund (dac) as a paradigm, we previously showed that the loss of PRE contacts induced a very specific gain-of-function phenotype in the adult fly leg, without affecting gene expression during early development²². While this suggests that PRE loops have repressive functions and contribute to stabilize gene silencing during development, another PRE at the homeobox gene locus that coincides with a loop anchor of TEs is involved in gene activation at the embryonic stage¹⁵. This suggests that chromatin loops involving PREs can mediate both gene activation and repression but many questions concerning PRE loop formation and their functions remain unsolved. It has not been defined whether PRE loops are spatially or temporally regulated and the effects of the loss of PRE contacts on the global 3D chromatin architecture of Polycomb TADs are unknown. Furthermore, the nature, timing and underlying molecular mechanisms of gene misexpression upon loss of a PRE loop remain to be studied.

To address these questions, we exploited the dac Polycomb domain by generating mutant fly lines in which we specifically interfered with PRE contacts and/or enhancer function and analyzed the effects on TAD architecture and gene expression during Drosophila development. Intriguingly, both PREs at the dac locus were identified as TEs in early embryogenesis¹⁵. Here, we show that the PRE loop constitutes a 3D chromatin scaffold of the Polycomb domain that forms independently of gene expression states and is present at all stages during fly development. Insertion of an insulator at various positions generates local insulation within Polycomb domains without creating a new TAD boundary. Intriguingly, the insulator blocks PRE looping, whereas an essential leg enhancer can bypass the insulator to activate the dac gene. Loss of PRE looping does not lead to widespread gene activation but correlates with the activation of the dac gene at a precise developmental stage and tissue. Importantly, the modulation of physical proximity between the leg enhancer and promoter is not a major determinant for gene activation. Lastly, we show that reduced PRE looping results in a loss of enhancer specificity, leading to ectopic activation of a neighboring gene. We propose that PRE loops form a topological scaffold structure within Polycomb domains that have a dual function; they restrict promoter–enhancer communication, while they also contribute to enhancer–promoter specificity.

Results

dac expression during development upon loss of PRE looping

To analyze the importance of PRE looping for the 3D organization of Polycomb domains and gene expression, we created three classes of clustered regularly interspaced short palindromic repeats (CRISPR)–Cas9 mutant flies associated with the dac gene locus (Fig. 1a). The first class comprised PRE deletion lines that affect both Polycomb group (PcG) recruitment and PRE looping²². The second category corresponded to the deletion of an enhancer driving dac expression in the leg. The third class consisted of inserting a gypsy insulator sequence between two PREs at various positions upstream or downstream of the leg enhancer.

**Fig. 1: Expression of *dac* gene during *Drosophila* development upon loss of PRE function and/or PRE looping.**

We previously showed that the disruption of PRE looping does not change the embryonic expression pattern of the dac gene. However, a very specific gain-of-function phenotype was observed in the adult fly: the appearance of extra sex comb (ESC) bristles on the second tarsal segment (TS2) of male flies on the first legs²². To determine the nature and timing of dac misexpression inducing this phenotype, we analyzed dac expression patterns during leg development in PRE deletion fly lines by performing immunostaining of third-instar larval imaginal leg discs and early pupal imaginal leg discs, when leg segmentation takes place. In larval leg discs, dac is expressed in a ring-like shape corresponding to the medial leg structures (trochanter, femur, tibia and TS1) and no notable changes in dac expression pattern can be observed at the larval stage upon mutation of PRE sequences (Fig. 1b).

At the early pupal stage, dac is normally expressed in the proximal leg part and TS1, whereas expression in TS2 is low and it is completely absent in the more distal segments (TS3–TS5) (Fig. 1c). Importantly, the expression of dac is significantly increased, specifically in TS2 of the developing leg in PRE mutant fly lines, whereas dac remains repressed in TS3–TS5 (Fig. 1c,d). No significant difference in global mRNA levels was observed by reverse transcription (RT)–qPCR analysis in pupal and larval imaginal leg discs (Extended Data Fig. 1a). This shows that PRE deletion leads to derepression of dac specifically in cells of the TS2 segment of pupal leg discs. Increased dac expression at this precise developmental stage and tissue is likely involved in transforming TS2 into TS1 identity, consistent with the induction of ectopic sex combs (normally found exclusively on the more proximal segment TS1) on the more distal segment TS2 (Extended Data Fig. 1b).

The PRE loop acts as a constitutive 3D chromatin scaffold

We hypothesized that changes in DAC expression, leading to the transformation of leg segment identity, may involve alterations in 3D chromatin architecture in mutant flies. To test this, we performed Hi-C experiments of third-instar larval imaginal leg discs and early pupal leg discs and compared them to Hi-C data in embryos (Fig. 2a). Interestingly, dac intra-TAD interactions decrease during development (Fig. 2b). This might reflect global decondensation of the domain correlating with an increase in dac-expressing cells during the larval and pupal stages (approximately 20% dac-expressing cells in late embryos versus 50% dac-expressing cells in leg discs). Importantly, the local contact enrichment of the PRE loop is present at similar levels from embryos to pupae (Fig. 2c). Furthermore, no additional chromatin loop or major change in intra-TAD structure or TAD borders was observed during these stages of development. This indicates that the PRE loop is a prominent chromatin feature of the dac TAD that persists at all stages of Drosophila development. Because the proportion of dac-expressing cells greatly increases during development, we hypothesized that the PRE loop within the dac TAD may not be the key element determining cell-type-specific expression.

**Fig. 2: PRE loop during *Drosophila* development and *dac*-expressing or *dac*-repressed cells.**

To test this model, we determined the conformation of the dac TAD in the different leg segments expressing or not expressing dac. For this, we turned to Hi-M, a multiplexed DNA fluorescence in situ hybridization (FISH) technology that captures chromatin conformations in single cells²⁶. We designed and amplified an oligopaint library tiling most of the dac TAD with different barcodes, reaching a mean resolution of 4 kb (Supplementary Table 1). This library was hybridized and each barcode was sequentially imaged in early pupal leg discs (Methods). Ensemble Hi-M maps obtained from pooled nuclei comprising TS1–TS4 confirmed the presence of a prominent long-range chromatin interaction between barcodes including and adjacent to the two PRE sequences (Fig. 2d). Hi-M maps obtained from different replicates were highly correlated (Extended Data Fig. 2b). As in Hi-C maps, the PRE loop includes additional neighboring chromatin regions, indicating that it involves an extensive set of chromatin interactions in the whole regions surrounding the PREs. Next, we analyzed the chromatin organization of the dac TAD in the different TSs, where the dac gene is highly expressed (TS1), weakly expressed (TS2) or repressed (TS3 and TS4). Interestingly, PRE interactions can be observed with similar frequency in all these segments (Fig. 2e). Taken together, these results show that the dac PRE loop forms with similar frequencies at different stages of development and within leg segments displaying different levels of dac expression. Thus, we conclude that the regulation of dac expression does not require changes in PRE looping.

A ring enhancer (RE) is necessary and sufficient for dac expression

Because the loss of PRE function induces activation of the dac gene exclusively during leg development, we searched for putative regulatory regions driving expression of the dac gene specifically in the leg. Interestingly, a well-conserved 567-bp sequence was previously shown to recapitulate dac expression in a ring-like shape in larval leg discs in transgenic reporter gene assays²⁷. Therefore, this regulatory region 20 kb downstream of the dac promoter (Fig. 1a) was called an RE.

Knowing that the RE is sufficient to recapitulate dac expression in reporter assays, we asked whether the RE is essential for regulating the correct dac expression pattern at the endogenous dac TAD. Therefore, we created a mutant fly line carrying a 1-kb deletion encompassing the RE sequence (ΔRE) (Fig. 1a). Heterozygous flies for the RE deletion were viable and did not show any morphological phenotype. In contrast, homozygous flies displayed short and deformed legs with fused segments, characteristic of dac loss of function (Fig. 3a). Strong loss of dac expression was observed by immunostaining experiments in imaginal leg discs and early pupal discs (Fig. 3b). Although dac expression was absent in the large majority of cells, a few random spots of cells expressing dac could be observed and morphological perturbations of the imaginal disc appeared at the early pupal stage during metamorphosis (Fig. 3b).

**Fig. 3: Phenotypic, transcriptional and 3D chromatin conformational consequences of the RE deletion.**

Together, these results show that the RE is the major CRE driving the expression of the dac gene in larval and early pupal leg discs and is both necessary and sufficient for dac activation in the leg disc.

Loss of dac expression and PRE looping

RE deletion with a concomitant loss of dac expression affects the formation and topology of the repressive H3K27me3 domain (PRE looping in particular).

Therefore, we first performed Hi-C experiments in the wild-type (WT) or ΔRE larval imaginal leg discs to determine the consequences of RE deletion on the PRE loop and global TAD architecture (Fig. 3c). Analysis of the Hi-C data and quantification of the interaction frequency of the PRE loop revealed that overall TAD structure, boundaries and PRE looping were not significantly affected upon deletion of the RE (Fig. 3d,e).

Next, we performed cleavage under targets and release using nuclease (CUT&RUN) experiments using H3K27me3 antibodies in WT or ΔRE leg imaginal discs (Fig. 3f). We did not observe any major changes in the distribution of the repressive H3K27me3 mark across the dac locus and quantitative analysis of H3K27me3 levels confirmed that the deposition of H3K27me3 across the TAD was not notably changed upon deletion of the RE. This indicates that the RE does not have a role in PRE function and the deposition of H3K27me3. Overall, these experiments show that the PRE loop is formed independently of dac expression states and the RE has no effect on PRE function and global TAD architecture.

Gypsy insertion induces insulation and reduces PRE looping

The identification and characterization of the RE as an essential CRE driving dac expression in the leg strongly suggest that derepression of dac in TS2 upon loss of PRE function is mediated by the RE. To disentangle the functional relationship of PRE looping, RE–promoter communication and dac expression, we generated a panel of mutant fly lines, where we inserted a gypsy insulator between the RE and the dac promoter and/or between the two PREs (Fig. 1a). The gypsy insulator element has the ability to reduce enhancer–promoter or PRE–PRE interactions when placed between these elements^28,29. In a previously characterized fly line (gypsy 1), the insulator was inserted upstream of the RE and was not located between the RE and dac promoter (gypsy 1 line, Fig. 1a). This insulator is not expected to interfere with the interaction of the RE and the dac promoter. We, therefore, created two lines where we inserted the gypsy insulator at different positions between the RE and the dac transcription start site (TSS) (gypsy 2 and gypsy 3 lines, Fig. 1a). We hypothesized that, in these lines, the gypsy element should interfere with both PRE looping and RE–promoter communication.

Quantitative chromatin immunoprecipitation (qChIP) experiments confirmed that all three gypsy insertions generated an ectopic Su(Hw)-binding site, indicating that the insulator sequence is functional (gypsy 2 and gypsy 3 lines, Extended Data Fig. 3a; Su(Hw) binding to gypsy 1 was analyzed in a previous study²²). Additionally, CUT&RUN experiments in larval imaginal leg discs showed that PRE-mediated deposition of H3K27me3 was not notably changed upon insertion of the gypsy insulator in all three gypsy lines, confirming that the recruitment of PcG complexes to PREs is not affected (Extended Data Fig. 3b,c).

To analyze the physical insulation activity of the gypsy element and its impact on PRE looping, we performed Hi-C experiments in larval imaginal leg discs of the three gypsy insertion lines (Fig. 4). Visual inspection of Hi-C contact maps revealed that no new TAD borders were formed around the gypsy insertion sites (Fig. 4a). A moderate insulation activity of the gypsy sequences could be visualized by plotting the maps of differential score enrichments of interactions in the WT line versus each of the gypsy mutant lines (Fig. 4b). Gypsy 1 insertion had the weakest effect on local insulation, whereas, in comparison to gypsy 2 and gypsy 3 lines, the WT had stronger contact enrichment in the region bypassing the gypsy insertion sites, indicating an insulation activity of gypsy on the surrounding chromatin. Accordingly, quantification of the insulation scores (ISs) at the gypsy insertion sites showed a significant increase in insulation at the gypsy 2 and gypsy 3 insertion sites, whereas no significant differences between the insulation profiles of the different conditions were observed in the region upstream of the PRE1 site (Extended Data Fig. 4a,b). Although gypsy 1 insertion showed no significant increase in local insulation in the Hi-C approach (Fig. 4c,d), we observed an insulation of the surrounding chromatin at the gypsy 1 insertion site insulation activities using 4C-seq experiments with viewpoints upstream or downstream of the gypsy insertion (Extended Data Fig. 5a,b). This suggests that, albeit with different strength, all three gypsy elements have insulation activity.

**Fig. 4: Consequences of gypsy insertions for the *dac* Polycomb domain architecture analyzed by Hi-C.**

Next, we analyzed how the physical insulation activity in the different gypsy lines affects the PRE loop. Quantification of the looping interactions between the two PRE regions revealed a significant reduction in PRE contacts upon gypsy insertions in each of the three mutant lines (Fig. 4e). Quantitative differences were still observed between the gypsy lines, with gypsy 1 insertion weakly reducing PRE contacts, whereas gypsy 2 and gypsy 3 insertions had stronger effects on PRE looping. Reduced PRE looping of the gypsy 1 line was further confirmed by 4C-seq experiments using the dac promoter as a viewpoint (Extended Data Fig. 5b).

Lastly, we analyzed changes in gypsy insulation activity and its effects on PRE looping during Drosophila development. For this, we exclusively used the gypsy 2 line and performed 4C-seq experiments at different developmental stages (embryos, larval leg discs and pupal leg discs). Using the dac promoter (PRE2) (Extended Data Fig. 6a) and the RE (Extended Data Fig. 6b) as viewpoints, we observed that gypsy insertion resulted in a similar physical insulation activity at all developmental stages and PRE looping was reduced at all developmental stages investigated (Extended Data Fig. 6a,b).

Altogether, these experiments indicated that gypsy insertion interferes with PRE looping during all stages of fly development investigated, although the position of insertion of the gypsy insulator affects the strength of the insulation effect.

Gypsy insulator does not block RE function

Next, we asked whether gypsy insertions affect enhancer–promoter interactions and change dac expression. We predicted that insertion of the gypsy insulator upstream of the RE (gypsy 1), which interferes with PRE looping, might induce dac expression in TS2, as is the case in the PRE deletion lines (Fig. 1). On the other hand, gypsy 2 and gypsy 3 insulators inserted between the RE and the dac promoter might block enhancer–promoter communication.

Intriguingly, insertion of the gypsy insulators between the RE and the dac promoter (gypsy 2 and gypsy 3) resulted in a significant increase in dac expression in TS2 of pupal leg discs (Fig. 5a,b), whereas dac activation upon gypsy insertion downstream of the RE (gypsy 1) was much weaker. Moreover, activation of dac in TS2 in the gypsy 2 and gypsy 3 lines induced the dac gain-of-function phenotype (ESC) on adult male fly legs (Fig. 5c). The penetrance of ESC in the gypsy 2 and gypsy 3 lines (about 25% and 50% of all male flies, respectively) was significantly stronger than the penetrance observed in the gypsy 1 line. Notably, there was a good correlation between the penetrance of ESC and the levels of overexpression of dac in TS2 (Fig. 5b,c), which in turn inversely correlated with the reduction in PRE contacts (Fig. 4e), suggesting that loss of PRE looping and dac activation in TS2 are functionally linked.

**Fig. 5: Transcriptional and phenotypic consequences of gypsy insulator insertions at the *dac* Polycomb domain.**

The absence of enhancer-blocking activity when the insulator was inserted between the RE and promoter is surprising, given the previously reported enhancer-blocking function of the gypsy element³⁰. One possible explanation could be that gypsy blocks the RE but, in this case, a shadow enhancer might take over and begin to induce dac expression.

To test whether the overexpression of dac upon gypsy insertions is induced by the RE, we deleted the RE in the presence of the gypsy 2 insertion (gypsy 2 + ΔRE line; Fig. 1a). Indeed, if the gypsy insertion induces dac overexpression independently of the RE, we would expect at least a partial rescue of the loss-of-function phenotype upon deletion of the RE (Fig. 3a). Instead, we observed that homozygous flies carrying gypsy 2 + ΔRE displayed the same strong crippled leg phenotype compared to the deletion of the RE alone, with a complete penetrance (Fig. 5d). Furthermore, RT–qPCR analysis showed that dac expression was reduced to the same extent in the gypsy 2 + ΔRE line compared to the ΔRE alone (Fig. 5e).

Altogether, these results indicate that the RE can bypass the gypsy insulator to activate dac expression irrespective of the genomic location of the gypsy insulator. Moreover, activation of the dac gene upon gypsy insertion and reduced PRE looping are strictly dependent on the presence of RE.

Unchanged enhancer–promoter proximity upon dac gene activation

Next, we tested whether the bypass of the insulator and activation of the dac gene involves changes in the physical proximity between enhancer–promoter sequences. For this, we performed Hi-M experiments in pupal leg discs in the gypsy 2 mutant, which allowed us to analyze RE–promoter distances specifically in the TS where dac is activated. Comparing the interaction profiles of gypsy 2 mutant cells to WT cells confirmed that the PRE loop is reduced in all TSs (TS1, TS2 or TS3 and TS4) independent of the dac expression status (Extended Data Fig. 7a). Furthermore, we observed increased short-range interactions upstream or downstream of the gypsy insulator, whereas long-range interactions between sequences upstream and downstream of the insulator insertion were reduced, consistent with the insulator activity of the gypsy sequence (Extended Data Fig. 7a,b).

Next, we compared the distances between the RE and the dac promoter in the TSs where dac is active (TS1), moderately expressed (TS2) or inactive (TS3 and TS4) in WT or gypsy 2 mutant flies. Therefore, we created virtual 4C plots derived from Hi-M experiments using the dac promoter as a viewpoint (Fig. 6a). Remarkably, despite the upregulation of dac in the gypsy mutant, the RE–dac promoter distance was not decreased but displayed a very modest increase between WT and the gypsy mutant in TS2 (≤10 nm). A similar weak increase in RE–promoter distances was observed in TSs where dac remains repressed (TS3 and TS4), indicating that this increase in RE–promoter distance was not sufficient for gene activation. We note that the RE–dac promoter distance changes were statistically significant for these segments (Wilcoxon two-sided rank test, Fig. 6a) but were, in all cases, comparable to the measurement error in the median distance as estimated by bootstrapping analysis (~8–25 nm; Methods). We, therefore, concluded that activation of dac expression in TS2 upon loss of PRE looping is not the consequence of large-scale changes in RE–promoter distances. Likewise, RE–dac promoter distances exhibited no significant changes when we compared TS1 (where dac is active) to TS3 and TS4 (where dac is completely repressed) in WT pupal leg discs (Fig. 6b). Importantly, the distance distributions across individual cells between PREs or enhancer–promoter elements showed no evidence for bimodality in all TSs including TS2, which comprises a mixed population of dac expressed and repressed cells (Extended Data Fig. 8a,b). This argues against the existence of different chromatin conformations within the examined TSs.

**Fig. 6: RE–*dac* promoter distances in TSs of early pupal leg discs in WT versus gypsy 2 flies determined by Hi-M.**

Together, these results indicate that increased physical proximity between the RE and dac promoter does not appear to be a major mechanism involved in dac gene activation.

The PRE loop contributes to enhancer–promoter specificity

Because the major structural effect of gypsy insertion is to reduce PRE looping at the dac domain, we asked whether other genes within the domain are affected by the loss of looping interactions. Therefore, we first performed RT–qPCR analysis at different developmental stages in mutant fly lines, where we interfered with either PRE function (double, ΔPRE2), PRE looping (gypsy 2) or enhancer function (ΔRE) (Fig. 7a). We found that the CG5888 gene was induced already at the larval stage upon loss of PRE contacts. The Idgf1 gene was also activated, whereas two other genes, Idgf2 and Idgf3, remained repressed at all developmental stages analyzed (Extended Data Fig. 9a). RNA FISH analysis confirmed the transcriptional activation of CG5888 upon the reduction in PRE looping in both gypsy 2 and gypsy 3 larval leg discs (Fig. 7b and Extended Data Fig. 9b,c). Interestingly, CG5888 was mainly overexpressed in cells that also expressed dac and where the RE was active (Fig. 7c), resulting in a similar ring-shaped expression. Importantly, expression of the CG5888 gene was not significantly changed upon deletion of the RE alone (Fig. 7b,d,e), indicating that CG5888 expression is not controlled by the RE in WT conditions. However, the ectopic expression of the CG5888 gene upon PRE looping reduction depends on the presence of the RE because CG5888 activation was lost when we deleted the RE in the presence of the gypsy 2 insertion (ΔRE + gypsy 2) (Fig. 7b,d,e). Importantly, the perturbation of the PRE loop did not lead to the activation of all genes within the dac TAD (Extended Data Fig. 9a) and was restricted to the region where the RE was active during leg development. This indicates that the loss of the PRE loop does not create a global permissive environment facilitating transcription per se but rather regulates RE specificity toward permissive promoters such as the CG5888 gene. Altogether, these results suggest that, in addition to TADs, which can restrict enhancer–promoter communication across TAD borders, PRE loops within TADs constitute an additional gene-regulatory layer by contributing to intra-TAD enhancer specificity (Extended Data Fig. 10a).

**Fig. 7: Consequences of gypsy insertions and/or RE deletion on *CG5888* and *dac* gene expression.**

Discussion

We showed that the dac PRE loop constitutes a topological chromatin structure that has a versatile function. On one hand, it can regulate enhancer–promoter communication in a manner specific to the developmental stage and tissue. On the other hand, the PRE loop contributes to enhancer–promoter specificity by restricting the enhancer activity to its specific target promoter (the dac gene), as shown by the RE-dependent illegitimate activation of the CG5888 gene upon insertion of a gypsy insulator between the two PREs.

Gypsy insulator reduces PRE looping but not enhancer function

Insulator elements are DNA sequences that act as chromatin boundaries and regulate interactions between genomic regulatory elements. The gypsy element is one of the best-characterized insulators. It contains three core components, Su(Hw), CP190 and Mod(mdg4), and the zinc finger protein CLAMP that promotes gypsy enhancer-blocking activity³¹.

At the dac TAD, the gypsy insulator sequence interferes with the interaction of the two PREs when inserted between them. However, gypsy insertion between the RE and the dac promoter does not block enhancer–promoter communication. Although surprising, these results are consistent with previous work showing that the Su(Hw) protein binds at thousands of sites throughout the genome yet is not systematically associated with the physical boundaries of gene units⁵. Although more than 20 enhancers have been shown to be blocked by the gypsy insulator (as discussed in a previous study³⁰ and references therein), this element does not necessarily establish an impermeable chromatin barrier^32,33. In all the enhancer–promoter pairs that were analyzed in these studies, each element of the pair was located close to the gypsy element and insulation might perhaps depend on the ability of the gypsy element to reduce short-range chromatin contacts. Furthermore, a recent genome-wide analysis suggested that gypsy-binding proteins do not generally act by blocking loop formation but rather by inducing local insulation at their binding sites³⁴.

In transgenic reporter assays, the gypsy insulator behaves as a chromatin border that is able to block spreading of H3K27me3 and prohibit contacts between a PRE and a distal promoter^29,35. In contrast, at the dac TAD, insertion of the same gypsy insulator between the two PREs does not interfere with the formation of the repressive H3K27me3 domain. This is in agreement with the presence of endogenous Su(Hw)-binding sites within the dac TAD that, at this genomic locus, do not act as classical chromatin domain borders and do not interfere with the deposition of H3K27me3. This indicates that the function and effect of the gypsy insulator on the physical interaction between CREs or the partitioning of chromatin domains are context dependent.

PRE contacts constitute a specific form of TE loops

A recent paper used the Micro-C technique to demonstrate that the genome is organized by insulator elements and so-called TEs in early Drosophila embryos¹⁵. These TEs correspond to ‘organizational elements’ that form chromatin loops to facilitate specific enhancer–promoter contacts for rapid gene activation.

Intriguingly TEs and PREs show several similarities. First, both PRE loop and TE loop disruptions have little effect on the overall structure of TADs, whereas they have an impact on enhancer–promoter communication. Secondly, both elements are frequently bound by the GAGA factor, which has been proposed to mediate chromatin loops³⁶. Lastly and most compellingly, we observed that 48% (109 of 225) of PREs identified in embryos coincided with TEs (Extended Data Fig. 10b), notably including the two PREs of the dac gene locus. We, therefore, propose that PRE loops actually constitute a specific form of TE loops.

In contrast to the dac PRE loop, which restricts enhancer–promoter communication in pupal leg discs, a chromatin loop between TEs (that also corresponds to PREs) at the scr locus was shown to promote enhancer–promoter contacts¹⁵. Another example of a TE loop that corresponds to a PRE loop involves the regulation of the cut (ct) gene. Importantly, a natural insertion of a gypsy retrotransposon between the two PREs and TEs of the ct gene locus leads to ct downregulation and loss of enhancer–promoter communication²⁸. Thus, chromatin loops involving PREs and TEs can clearly mediate both gene activation and repression. Intriguingly deletion of both looping anchor points of the dac PRE loop resulted in reduced dac gene expression in embryos²². Although this reduced expression does not result in developmental defects, this suggests that even the same PRE or TE loop can have different functions in gene activation and repression depending on the developmental stage or tissue. These antipodal functions of PRE and TE loops might be mediated by the presence of developmental or stage-specific factors. Interestingly, a recent study analyzed chromatin loops of paralogous gene pairs, proposing an additional function of chromatin loops in the fine-tuning of coordinated expression levels of genes with related function³⁷. As is the case for the dac PRE loop, the same chromatin loop can have different functions in the regulation of gene expression³⁷.

Together, these data suggest that PRE and TE loops constitute a chromatin scaffold structure that is used to regulate enhancer–promoter communication positively or negatively, depending on the gene locus, the developmental stage and the specific tissue.

PRE looping as a regulator of enhancer–promoter communication

Loss of PcG binding to the PREs is unlikely to be the major mechanism leading to dac gene activation upon loss of PRE looping. We previously observed that deletion of both PREs, with concomitant loss of PcG binding, was not sufficient to globally activate dac expression but only resulted in the same tissue-specific activation of the dac gene as the insertion of the gypsy insulator. Moreover, PcG proteins were shown to remain associated with PREs when their target genes are active^38,39. In contrast, activation of the dac gene in TS2 upon loss of PRE looping is strictly dependent on the RE, indicating that gene activation involves changes in enhancer–promoter communication.

A popular model of enhancer–promoter communication proposes chromatin looping as a mechanism to bring the two regulatory regions in close proximity⁴⁰ and experimentally forced enhancer–promoter looping contributes to gene activation⁴¹. By applying Hi-M to detect chromatin 3D organization in single nuclei⁴², including loops at distances similar to that separating the RE from the dac TSS, we were unable to detect increased contact frequency between the RE and the dac promoter compared to surrounding regions. In addition, no significant differences in contact frequencies of the dac promoter and RE were observed in repressed versus active cells, or upon loss of PRE looping, suggesting that no stable contact is needed for dac transcription. This agrees with a previous report showing no differences between enhancer–promoter pairs in active or inactive transcriptional states during early Drosophila development⁴². In addition, live-cell imaging analyses challenge the idea of stable loops as a general mechanism for all enhancer–promoter communication^43,44. Lastly, increased enhancer–promoter distance has been reported to accompany Shh gene activation during neural differentiation⁴⁵, supporting evidence that enhancers can act at a distance.

Several ‘action-at-a-distance’ models have been proposed to explain the functional interaction of distant enhancers with their promoters⁴⁶, including liquid–liquid phase separation, where enhancers function as binding surfaces that concentrate transcriptional activators in ‘transcriptional condensates’, allowing them to induce gene expression even when promoters are not in absolute proximity. Alternatively, in the transcription factor activity gradient model, enhancer-bound coactivators can activate target promoters within a ‘permissive range’. It is conceivable that the PRE loop might act as a crucial scaffold to create a particular chromatin environment or a ‘permissive range’, thereby regulating activation of the dac gene and enhancer specificity.

It is worth mentioning that the RE at the dac locus is not associated with the previously well-established molecular signatures of active enhancers (such as H3K4me1 and H3K27Ac enrichment) in embryos or larval disc tissues⁴⁷. This surprising lack of histone marks at an essential enhancer might indicate that other such enhancers with an important function but undetectable epigenomic signature exist in the genome. In the future, it will be interesting to study whether chromatin looping or contact-independent mechanisms are differently involved in gene activation for different classes of enhancer elements.

Methods

Fly work and generation of mutant flies by CRISPR–Cas9 genome engineering

All flies were raised on standard corn meal yeast extract medium at 25 °C. CRISPR–Cas9 mutant fly lines double, ΔPRE1, ΔPRE2 and gypsy 1 were described in a previous study²². Sequences of guide RNAs (gRNAs) used to create fly lines gypsy 2, gypsy 3, ΔRE and gypsy 2 + ΔRE are described in Supplementary Table 2. Sense and antisense oligonucleotides were annealed and phosphorylated by the T4 polynucleotide kinase (New England Biolabs, M0201S) before being inserted inside a pCFD3 plasmid (Addgene, 49410) previously digested by BbsI (NEB, R0539S). To create the pHD-dsRED donor plasmid (Addgene) containing a removable (floxed) 3XP3–dsRED construct flanked by loxP sites and DNA fragments having homology to the target regions (homology arms) serving as a template for homology-directed repair, 1.5-kb genomic DNA fragments were amplified by PCR (Supplementary Table 2) and inserted into the pHD-dsRED plasmid using Gibson assembly (kit NEBuilder; New England Biolabs, E2621S).

The gypsy insulator was amplified from the plasmid (Gy)w(Gy) described in a previous study⁴⁸ and introduced into the donor plasmid cut by SpeI and BglII using Gibson cloning (Supplementary Table 2). To generate mutant fly lines, gRNA-containing pCFD3 and pHD-dsRED donor plasmids were injected into flies expressing Cas9 in the germline (vas-Cas9(X) RFP⁻; Bloomington, 55821). Injections and dsRED screening were performed by BestGene (https://www.thebestgene.com/). To remove the dsRED reporter construct, mutant flies were crossed with a fly line expressing CRE recombinase (Bloomington, 34516). To generate the gypsy 2 + ΔRE mutant line, gRNAs targeting the RE and corresponding donor plasmid were injected into gypsy 2 mutant lines previously generated and expressing Cas9 (vas-Cas9(III)). Coordinates and sequences of deleted regions can be found in Supplementary Table 3. Genotypes of mutant fly lines were confirmed by PCR genotyping and sequencing analysis of the mutated region.

Immunostaining experiments

For immunostaining, third-instar imaginal leg discs were dissected at room temperature in sterile Schneider medium. Pupae were selected at the very beginning of pupation, where pupae can be recognized by their white color (a pupal stage that lasts 1 h), and were dissected 3.5–4 h later. The discs were then fixed for 20 min in 4% formaldehyde and were permeabilized for 1 h in PBS + 0.5% Triton (for larval leg discs) or 0.8% Triton (for pupal leg discs). The samples were then incubated for 1 h in 3% BSA PBTr (1× PBS + 0.5% Triton X-100). DAC primary antibody was diluted 1:400 (DSHB mAbdac1-1) in 1% BSA PBTr and incubated overnight at 4 °C on a rotating wheel. The leg discs were washed in PBTr before adding the secondary antibody at 1:1,000 dilution (Thermo Fisher Scientific, A-31571) and incubated for 2 h at room temperature on a rotating wheel. Finally, the discs were extensively washed in PBTr. The proximal segments of the leg discs were removed by dissection to only keep the TSs, which were subsequently mounted on microscope slides using ProLong Gold reagent (Invitrogen, P36930). The different images were acquired on a Zeiss axioimager Z2 Apoptome Leica SP8 confocal microscope using the same settings for all mutant lines and analyzed using Fiji software.

RNA FISH experiments

RNA FISH probes were prepared with an RNA FISH probe kit (Thermo Fisher Scientific, F32956) from DNA probes amplified with the primers described in Supplementary Table 2. Third-instar imaginal leg discs were quickly dissected in Schneider medium. Pupae were selected at the very beginning of pupation, where pupae can be recognized by their white color (a pupal stage that lasts 1 h), and were dissected 3.5–4 h later. The discs were then fixed with 4% formaldehyde before being permeabilized with PBTr for 4 h. Subsequently, discs were incubated for 10 min with 50% PBT (PBS + 1% Triton), 50% hybridization solution (50% formamide, 5× saline–sodium citrate (SSC), 100 g ml⁻¹ fragmented salmon testes DNA, 50 g ml⁻¹ heparin and 0.2% Tween-20) at room temperature. The samples were incubated for 45 min and then 1 h in hybridization solution at 55 °C. In parallel, a previously tested optimal concentration of labeled probe was diluted in 50 µl of hybridization solution, heated for 2 min at 85 °C and chilled on ice to denature RNA secondary structures. The discs were then incubated overnight with 50 µl of probe solution at 55 °C. The day after, the samples were washed three times at 55 °C with hybridization solution and twice with PBT. The proximal segments of the leg discs are removed by dissection to only keep the TSs, which were mounted on microscope slides using ProLong Gold reagent (Invitrogen, P36930). Images were acquired on a Zeiss axioimager Z2 APopoteme Leca SP8 confocal microscope using the same settings for all mutant lines and analyzed using Fiji software.

Hi-C experiments

Hi-C experiments were performed using the EpiTect Hi-C Kit (Qiagen, 59971). All Hi-C experiments were performed in two or three independent experiments using 50 third-instar imaginal leg discs or early pupal discs. Briefly, discs were homogenized and fixed in activated buffer T and 2% formaldehyde using tissue masher tubes (Biomasher II (EOG-sterilized), 320103). Tissue was digested by adding 25 μl of collagenase I and II (40 mg ml⁻¹) for 1 h at 37 °C. Samples were centrifuged and supernatant was carefully aspirated, leaving ~250 μl of solution in the tube. Then, 250 μl of QIAseq beads equilibrated to room temperature were added to bind nuclei to the beads and all subsequent reactions were performed on the beads according to the manufacturer’s protocol. Libraries were sequenced at BGI (https://www.bgi.com/) with 150-bp paired-end reads (approximately 400 million reads per replicate).

Hi-C analysis

Raw data from Hi-C sequencing were processed using the ‘scHiC2’ pipeline. Sequencing statistics are summarized in Supplementary Table 4. Valid interactions were stored in a database using the ‘misha’ R package (https://github.com/msauria/misha-package). Extracting the valid interactions from the misha database, the ‘shaman’ R package (https://bitbucket.org/tanaylab/shaman) was used for computing the Hi-C expected models, Hi-C scores with parameters k = 250 and k_exp = 500 (Figs. 2a, 3c and 4a) and differential Hi-C interaction scores with parameters k = 250 and k_exp = 250, with the compared datasets downsampled to have the same number of valid pairs in chromosome 2L (chr2L) for each comparison (Fig. 4b). Specifically, Hi-C scores quantify the contact enrichment (positive values) or depletion (negative values) of each bin of the map with respect to a statistical model used to evaluate the expected number of counts. To generate this expected model, we randomized the observed Hi-C contacts using a Markov chain Monte Carlo-like approach per chromosome⁴⁹. Shuffling was conducted such that the marginal coverage and decay of the number of observed contacts with the genomic distance were preserved but any features of genome organization (for example, TADs or loops) were not. These expected maps were generated for each biological replicate separately and contained twice the number of observed cis contacts. Next, the score for each contact in the observed contact matrix was calculated using the k nearest neighbors (kNN) strategy⁴⁹. In brief, the distributions of two-dimensional Euclidean distances between the observed contact and its nearest k_exp neighbors in the pooled observed and pooled expected (per cell type) data were compared, using Kolmogorov–Smirnov D statistics to visualize positive (higher density in observed data) and negative (lower density in observed data) enrichments. These D scores were then used for visualization (using a scale from −100 to +100) and are referred to as Hi-C scores in the text. Accordingly, the color scale of the Hi-C scores comprises both positive and negative values. When computing the differential Hi-C scores maps of Fig. 4b, the reference dataset was used as the expected model.

For each condition, the Hi-C interaction quantifications at the dac PRE loop (Figs. 2c, 3d and 4e) were performed by considering the Hi-C scores between two regions of 6 kb, chr2L:16419514–16425515 and chr2L:16482929–16488930), including the PRE1 and PRE2, respectively (Supplementary Table 3). The distributions of Hi-C scores (Figs. 2c, 3d and 4e) are represented as box plots showing the median (central line), the 75th and 25th percentiles (box limits) and 1.5 × the interquartile range (IQR; whiskers). Each of the comparisons of the Hi-C interaction quantifications at the dac PRE loop was performed between a reference condition (embryo in Fig. 2c and WT larvae in Figs. 3d and 4e) and each of the other conditions present in the same figure. An unpaired two-sided Wilcoxon statistical test (with the null hypothesis that the true median shift is equal to zero and assuming that the two variables are not normally distributed) was used to estimate the reported P values. The annotation of the Polycomb-associated TADs in chr2L from a previous study⁵ was used to compute the number of Hi-C interactions within the PcG TAD, which were then normalized by the total number of valid pairs at the corresponding developmental stage (embryo, larvae or pupae). The distributions of these interaction frequencies are shown in the violin and box plots of Fig. 2b as the log₂ ratios of embryo over the larval and pupal leg discs. The box plots show the median (central line), the 75th and 25th percentiles (box limits) and 1.5 × IQR (whiskers). An unpaired two-sided Wilcoxon statistical test (with the null hypothesis that the true median shift is equal to zero and assuming that the two variables are not normally distributed) was used to estimate the reported P values. The ISs⁵⁰ were computed on the observed Hi-C datasets binned at 2-kb resolution with windows of 100, 150, 200, 250 and 300 kb, resulting in five values per bin, and were stored in the misha database using an in-house R script. The mean and s.d. for each of the 2-kb bins were computed for the plots in Figs. 3e and 4c. The quantification of the ISs at gypsy insertions and R0–R12 regions was performed by applying a pairwise statistical comparison of the five IS quantifications per 2-kb bin. The P values in Fig. 4d and Extended Data Fig. 4b resulted from a Welch t-test (with the null hypothesis that the true difference in means is equal to zero and assuming that the variances of the samples are not equal) between the WT condition and each of the gypsy mutants at the corresponding locus. All plots of Hi-C maps (Figs. 2a, 3c and 4a,b), Hi-C interaction score comparisons (Figs. 2c, 3d and 4e), IS profiles (Figs. 3e and 4c) and P values of IS comparisons (Fig. 4d) were obtained with in-house R scripts.

Hi-M library preparation

The oligopaint library covering the dac region consists of 52-mer sequences with genome homology ordered from CustomArray. These sequences were obtained from the oligopaint public database (http://genetics.med.harvard.edu/oligopaints). From the initial design of the library, we selected 20-mers with an average probe density of 9–17 probes per kb. Each barcode contained 45 probes covering 3.8 kb on average (Supplementary Table 1). Each oligo was composed of five different regions: (1) a 21-nt forward universal priming region for library amplification; (2) two 20-nt readout regions separated by an A for barcoding; (3) a 42-nt genome homology region; (4) a duplication of one 20-nt readout region; and (5) a 21-nt reverse universal priming region.

The procedure for oligopaint library amplification was previously described^26,42,51,52. It consists of seven steps: (1) an emulsion PCR (emPCR) to extract the dac library from the oligonucleotide pool using universal primers; (2) a limited-cycle PCR performed on the emPCR to identify the most efficient amplification cycle; (3) a large-scale PCR with a T7 promoter on the reverse primer; (4) an in vitro T7 transcription; (5) RT to transform RNAs into single-stranded DNA (ssDNA); (6) an alkaline hydrolysis for the removal of the intermediate RNA; and (7) ssDNA purification and concentration.

Each barcode is unique and specific to an adaptor oligo. The adaptor oligo serves as a bridge between the readout region and an Alexa Fluor 647-labeled secondary oligonucleotide. The fluorescently labeled part of the secondary probe is attached by a disulfide leakage that can be cleaved (chemical bleaching) during the sequential imaging of FISH probes⁵¹. For the fiducial, we used an adaptor oligo complementary to the reverse primer of the library and specific to a secondary probe bound to a noncleavable rhodamine red fluorophore. Adaptors and fluorescently labeled secondary probes were synthesized and purchased from Integrated DNA Technologies.

Hi-M library hybridization

Pupae were collected at the beginning of pupation (white pupae) and dissected 3.5–4 h later. The dissected leg disc were fixed with 4% formaldehyde before being permeabilized with PBTr for 4 h. The discs were then progressively washed in four different concentrations of Triton and pHM (2× SSC and 0.1 M NaH₂PO₄ pH 7; 20%, 50%, 80% and 100%) for 20 min in each buffer at room temperature on a rotating wheel. Then, the discs were incubated overnight in 225 pmol of the library diluted in 30 µl of FISH hybridization buffer (FHB; 50% formamide, 2× SSC, 0.5 mg ml⁻¹ salmon sperm DNA and 10% dextran sulfate). The probes and the discs in pHM were heated at 80 °C. The incubation of the leg discs in the FHB + probe buffer was performed in a PCR machine from 80 °C to 37 °C with a temperature decrease of 1 °C every 10 min. The next day, discs were washed two times with 50% formamide, 2× SSC and 0.3% CHAPS and sequentially washed with four different concentrations of formamide and PBT (40%, 30%, 20% and 10% formamide) for 20 min per buffer on a rotating wheel. Finally, the discs were washed with PBT, fixed with 4% formaldehyde in PBS, washed with PBS and stored at 4 °C.

Hi-M imaging system

Hi-M experiments were performed with a homemade wide-field and epifluorescence microscope. This setup includes a rapid automated modular microscope (Applied Scientific Instrumentation) coupled with a microfluidic device as previously described^26,42. The microscope and fluidics system were controlled using Qudi-HiM (our homemade hardware control package)⁵³. The fluidics system permitted the automated and sequential hybridizations of the probes. The solutions were delivered to the sample by a combination of three eight-way valves (HVXM 8-5, Hamilton), a negative pressure pump (MFCS-EZ, Fluigent) and an FCS2 flow chamber (Bioptechs). The excitation was performed by three different lasers: 405 nm (Obis 405, 100 mW; Coherent), 561 nm (Sapphire 561 LP, 150 mW; Coherent) and 642 nm (VFL-0-1000-642-OEM1, 1 W; MPB communications). The fluorescence was collected through a Nikon APO ×60 (1.2 numerical aperture) water-immersion objective lens mounted on a closed-loop piezoelectric stage (Nano-F100, Mad City Labs). Images were acquired using a scientific complementary metal–oxide–semiconductor camera (ORCA Flash 4.0 V3, Hamamatsu) with an effective optical pixel size of 106 nm. To correct axial drift in real time, we used a homemade autofocus system composed of a 785-nm laser (OBIS 785, 100 mW; Coherent) and an infrared-sensitive camera (DCC1545M, Thorlabs).

Acquisition of Hi-M datasets

The proximal part of the pupal leg discs was removed by dissection to only keep the TSs. About 15–20 TSs were aligned on a 2% agar–PBS pad and then attached onto a 40-mm round coverslip previously functionalized with trimethoxysilane and 10% poly(l-lysine). The slide was then mounted onto the flow chamber. Pupal leg discs were first incubated with the fiducial adaptor (25 nM of the adaptor specific to the reverse primer, 2× SSC and 40% v/v formamide) for 20 min and then washed with a washing buffer solution (2× SSC and 40% v/v formamide) for 10 min. To complete the hybridization of the fiducial, we did a second round of incubation with the appropriate secondary oligo (25 nM of rhodamine-red-labeled probe, 2× SSC and 40% v/v formamide) for 20 min and washed again for 10 min with the washing buffer solution. After a 5-min wash with 2× SSC, we proceeded with nuclear staining with 0.5 µg ml⁻¹ of DAPI in PBS for 20 min. After another 5-min wash with 2× SSC, the imaging buffer (1× PBS, 5% glucose, 0.5 mg ml⁻¹ glucose oxidase and 0.05 mg ml⁻¹ catalase) was injected to limit fiducial photobleaching during the acquisition. An image stack (200 µm × 200 µm region of interest) was acquired for each of the 10–15 pupal leg discs. The DAPI and the fiducial were sequentially imaged (using 405-nm and 561-nm lasers) with a z step size of 250 nm for a total range of 17.5 µm.

Next, adaptor oligos and the secondary probe were sequentially hybridized, acquired and photobleached to image the whole dac oligopaint library. The following steps were performed for each of the 22 barcodes: (1) adaptor (40 nM of adaptor oligonucleotide, 2× SSC and 40% v/v formamide) injection and incubation for 10 min; (2) imaging probe (40 nM secondary probe, 2× SSC and 40% v/v formamide) injection and incubation for 10 min; (3) 10-min wash with washing buffer solution; (4) 5-min wash with 2× SSC; (5) imaging buffer injection and sequential acquisition of fiducial and barcode with 561-nm and 642-nm lasers; (6) chemical bleaching (2× SCC and 50 mM TCEP) of the imaging probe; and (7) 5-min wash with 2× SSC before a new cycle of hybridization.

Image processing and Hi-M analysis

Raw TIFF images were deconvolved using Huygens Professional 21.04 (Scientific Volume Imaging, https://svi.nl). Hi-M analysis was performed using pyHiM, a homemade analysis pipeline (https://pyhim.readthedocs.io/en/latest/)⁵⁴, as previously described⁵⁵. First, images were z-projected by applying either the sum for DAPI channels or the maximum intensity for the barcodes and fiducial. For each cycle of hybridization, fiducial images were used to register the corresponding barcode image using global and local registration methods. Barcodes and fiducials were segmented in 3D using a neural network, followed by 3D localization of the center of each barcode mask⁵⁴. The fiducial oligo bound to the universal priming regions, thus labeling the entire dac locus. Therefore, we built chromatin traces by combining the DNA FISH spots colocalizing within single fiducial masks. DAPI images were used to manually segment the different TSs (TS1, TS2, TS3, TS4 or TS5). Pairwise distance (PWD) matrices were calculated for each single chromatin trace. From a list of PWD maps, we calculated the proximity frequencies as the number of chromatin traces in which PWDs were within 250 nm, normalized by the number of chromatin traces containing both barcodes. Hi-M maps of the WT condition were generated from 51,622 total traces from 48 pupal leg discs from two independent biological replicates. Hi-M maps of the Gypsy 2 mutant were produced from 63,458 total traces of 51 pupal leg discs from two independent biological replicates. Hi-M matrices were generated for all the TSs combined (TS1, TS2, TS3, TS4 and TS5), as well as for TS1, TS2 and TS3 + TS4. Each trace contained at least 12% of the barcodes. Virtual 4C figures were obtained by plotting the PWDs between the anchored barcode or viewpoint with the remainder of the barcodes of an Hi-M matrix.

Wilcoxon two-sided rank tests between the PWD distributions of the barcodes containing the RE and the dac promoter were performed to test the hypothesis that two independent samples (for example, WT and gypsy mutant) were drawn from the same distribution. P values < 0.05 were considered significant to reject the hypothesis (that is, a 5% significance level).

We estimated the error in the measurement of the median RE–dac promoter distance by performing bootstrapping analysis. For this, we performed 1,000 bootstrapping cycles drawn from the experimental distribution of PWDs to estimate the s.d. in the determination of the median distance. The errors were between 8 and 25 nm for the WT condition.

4C-seq experiments

For 4C, either about 3,000 embryos were collected or 300 third-instar imaginal leg discs were dissected, homogenized and fixed in 2% formaldehyde diluted in nuclear permeabilization (NP) buffer (15 mM HEPES pH 7.6, 60 mM KCl, 15 mM NaCl, 4 mM MgCl₂, 0.1% Triton X-100, 0.5 mM DTT and 1× protease inhibitors (complete EDTA-free tablets; Roche, 11 873 580 001)) for 10 min at room temperature. Fixation was stopped by adding 2 M glycine for 5 min. The samples were then washed once in NP buffer and twice in 1.25× NEB3 buffer and the pellet of fixed cells was frozen in liquid nitrogen and conserved at −80 °C.

The chromatin pellet was then layered with 500 μl of 1.25× DpnII buffer without resuspension and centrifuged. The pellet was resuspended in 250 μl of 1.25× DpnII buffer. Then, 10 μl of 10% SDS was added and incubated for 20 min at 65 °C and 40 min at 37 °C. Chromatin was then split into 250-μl 1.25× DpnII buffer aliquots of 5–6 × 10⁶ cells and incubated for 1 h at 37 °C with 3.3% Triton X (final concentration). Samples were digested with 500 units of DpnII overnight. The day after, DpnII enzyme was inactivated by heating the samples at 65 °C for 20 min. The fragments were then ligated for 5 h at 16 °C with T4 ligase (2,000 units per µl) and digested overnight with proteinase K at 65 °C. The day after, RNA was degraded by RNAse A solution for 1 h at 37 °C. DNA was purified with Ampure beads without size selection and digested overnight with NlaIII enzyme. The next day, the DNA fragments were circularized by overnight ligation with T4 ligase (2,000 units per µl) in a large buffer volume. Next, circularized DNA was purified by Ampure beads without size selection. Lastly, 4C PCR was performed with the primers described in Supplementary Table 2. The amplified DNA was purified with Ampure beads. The sequencing libraries were produced with an Illumina kit (Illumina, 20015964). Sequencing (paired-end sequencing with 150-bp reads; approximately 4 Gb per sample) was performed by Novogene (https://en.novogene.com/).

4C-seq processing and analysis

Using a custom-made python script, FASTQ sequencing files were split using 4C primer sequences to obtain individual FASTQ files only containing reads from a single viewpoint per genotype and tissue type. Accordingly, the reads were trimmed to remove viewpoint sequences up to the restriction sites. Subsequently, the trimmed reads were aligned against the DM6 reference assembly using Bowtie⁵⁶ with the parameters ‘-a -v 0 -m 1’ (no mismatches and no multiple alignments allowed). The number of successfully aligned reads can be found in Supplementary Table 5. The aligned reads were mapped to restriction fragments and genomic bins of 1 kb using HiCdat⁵⁷ to obtain tabular files describing the number of reads (that is, contact frequencies) for a given fragment or genomic bin. All subsequent analysis steps were conducted using R. Depending on the 4C samples genotypes and viewpoints, contact frequencies arising from the viewpoint (±4 bins) and contact frequencies mapping to genotype-specific deletions were masked by setting them to zero (Supplementary Table 3). Then, data from individual samples were normalized for differing overall library size (counts per million).

To analyze differences between different genotypes, t-tests using triplicate data per genotype were performed for each 1-kb genomic bin along the region of interest (chr2L:1630000–16600000). No multiple-testing correction was performed. Subsequently, the differences of the average of triplicates were plotted and genomic bins that exhibited P values < 0.1 were highlighted.

qRT–PCR experiments

Embryos were collected in a 16–20-h developmental time window. Third-instar imaginal leg discs or early pupal leg discs (3.5–4 hs after pupation) were quickly dissected (<30 min) in Schneider medium and transferred into Trizol. RNA was extracted using Trizol reagent and purified using an RNA clean and concentrator kit (Zymo Research, R1015) following the instructions and using DNAse I (Qiagen, 79254). Then, 250 ng of purified RNA was used for RT using the Maxima first-strand complementary DNA synthesis kit for RT–qPCR with dsDNase (Thermo Fisher Scientific, K1671) following the manufacturer’s recommendations. Finally, quantification of the RT product was performed on a LightCycler 480 (Roche) with the primers listed in Supplementary Table 2. Data analysis was performed on LightCycler software. Expression levels were normalized to the housekeeping gene RP49.

qCHIP experiments

qChIP experiments were performed as described in a previous study⁵⁸ with minor modifications. Chromatin was sonicated using a Bioruptor Pico (Diagenode) for 7 min (30 s on, 30 s off). Su(HW) antibody was diluted 1:100 for the immunoprecipitation. After decrosslinking, DNA was purified using MicroChIP DiaPure columns from Diagenode. Enrichment of DNA fragments was analyzed using a real-time PCR LightCycler 480 (Roche). The primers used are indicated in Supplementary Table 2.

CUT&RUN experiments

CUT&RUN experiments were performed as described in the literature⁵⁹ with minor modifications. A total of 50 eye discs were dissected in Schneider medium, centrifuged for 3 min at 700g and washed twice with Wash+ buffer before the addition of concanavalin A-coated beads. MNase digestion (pAG-MNase Enzyme from Cell Signaling) was performed for 30 min on ice. After proteinase K digestion, DNA was recovered using SPRIselect beads and eluted in 50 μl of Tris-EDTA. DNA libraries for sequencing were prepared using the NEBNext Ultra II DNA library prep kit for Illumina. Sequencing (paired-end sequencing with 150-bp reads; approximately 2 Gb per sample) was performed by Novogene (https://en.novogene.com/). H3K27me3 antibody (Active Motif, 39155) was diluted 1:100. IgG antibody (1:100, Cell Signaling Technology, 2729S) was used as a control.

CUT&RUN analysis

The quality of the reads was assessed using FastQC. FASTQ files were aligned to the D. melanogaster reference genome dm6 using Bowtie 2 (version 2.4.2)⁶⁰ with the following parameters: ‘--local --very-sensitive-local --no-unal --no-mixed --no-discordant --phred33 -I 10 -X 700’. SAM files were compressed into BAM files using SAMtools (version 1.16.1) and reads with low mapping quality (Phred score < 30) were discarded. Duplicate reads were removed using Sambamba markdup (version 1.0.0)⁶¹ with the following parameters: ‘-r --hash-table-size 500000 --overflow-list-size 500000’. For visualization, replicates were merged using SAMtools ‘merge’ with default parameters and reads per kilobase per million mapped reads (RPKM)-normalized bigWig binary files were generated using the bamCoverage (version 3.5.5) function from deepTools2 (ref. ⁶²) with the following parameters: ‘--normalizeUsing RPKM --ignoreDuplicates -e 0 -bs 10’. Genome browser plots were generated using the pyGenomeTracks package (version 3.8)⁶³. The 131 Drosophila Polycomb domains²² were used for differential enrichment analysis using the DESeq2 method from the ‘DiffBind’ R package (version 3.12.0). Differential quantification results of H3K27me3 levels within Polycomb domains are summarized in Supplementary Table 6.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

The sequencing data reported in this paper were deposited to the Gene Expression Omnibus under accession number GSE247377. Source data are provided with this paper.

Code availability

All original code was deposited on GitHub (https://github.com/cavallifly/Denaud_et_al_NatStructMolBiol_2024) and is publicly available as of the date of publication. Any additional information required to reanalyze the data reported in this paper is available from the corresponding author upon request.

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Acknowledgements

We are grateful to V. Corces (Department of Human Genetics, Emory University School of Medicine, Atlanta) for the anti-Su(Hw) antibody. We would like to thank the MRI and Drosophila facilities (BioCampus Montpellier), CNRS, INSERM and the University of Montpellier. This work was supported by grants from the European Research Council (Advanced Grant 3DEpi, to G.C.), the European CHROMDESIGN ITN project (Marie Skłodowska-Curie grant agreement No 813327, to G.C.), the European E-RARE NEURO DISEASES grant ‘IMPACT’ from the Agence Nationale de la Recherche (PLASMADIFF3D, grant no. ANR-18-CE15-0010, and LIVCHROM, grant no. ANR-21-CE45-0011, to G.C.), the Fondation ARC (EpiMM3D, to G.C.), the Fondation pour la Recherche Médicale (EQU202303016280, to G.C.), the MSD Avenir Foundation (Project GENE-IGH) and the French National Cancer Institute (INCa, PIT-MM grant no. INCA-PLBIO18-362, to G.C.). This project was also funded by the European Union’s Horizon 2020 Research and Innovation Program (grant no. 724429, to M.N.), the Bettencourt-Schueller Foundation (prize ‘Coup d’élan pour la recherche Française’, to M.N.), France-BioImaging (grant no. ANR-10-INBS-04, ‘Investments for the Future’, to M.N.) and the Drosophila facility (BioCampus Montpellier, CNRS, INSERM and the University of Montpellier). The funders had no role in study design, data collection and analysis, decision to publish or preparation of the manuscript.

Author information

These authors contributed equally: Marion Bardou, Giorgio-Lucio Papadopoulos.

Authors and Affiliations

Institute of Human Genetics, UMR9002 CNRS, University of Montpellier, Montpellier, France
Sandrine Denaud, Giorgio-Lucio Papadopoulos, Marco Di Stefano, Gonzalo Sabarís, Bernd Schuettengruber & Giacomo Cavalli
Centre de Biologie Structurale, IUMR5048 CNRS, INSERM U1054, University of Montpellier, Montpellier, France
Marion Bardou & Marcelo Nollmann
Department of Plant and Microbial Biology and Zurich-Basel Plant Science Center, University of Zurich, Zurich, Switzerland
Stefan Grob

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Sandrine Denaud
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Marion Bardou
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Giorgio-Lucio Papadopoulos
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Stefan Grob
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Marco Di Stefano
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Gonzalo Sabarís
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Marcelo Nollmann
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Bernd Schuettengruber
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Giacomo Cavalli
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Contributions

G.C. and B.S. conceptualized the study. S.D., M.B. and B.S. performed the experiments. S.G. performed the bioinformatic analysis of 4C experiments. M.N. and M.B. conceptualized and designed the Hi-M experiments. M.B. performed the image acquisition and data analysis. G.P. and M.D.S. performed the bioinformatic analysis of Hi-C experiments. G.S. performed the analysis of CUT&RUN data. B.S. and G.C. wrote the manuscript. B.S., C.G. and M.N. edited the manuscript.

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Correspondence to Bernd Schuettengruber or Giacomo Cavalli.

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Nature Structural & Molecular Biology thanks Robert Illingworth, Son Nguyen and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available. Primary Handling Editors: Carolina Perdigoto, Jean Nakhle and Dimitris Typas, in collaboration with the Nature Structural & Molecular Biology team.

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Extended data

Extended Data Fig. 1 Dac Gain of function phenotype upon loss of PRE function.

(a) RT-qPCR analysis in the indicated fly lines and developmental stages using primers specific for dac gene. The fold change between the indicated mutant and wild type (control) line is shown. Three independent experiments have been performed. Data are presented as the mean values ± s.d (error bars) of replicates. (b) Representative examples of WT or PRE mutant 1^st leg male flies. The dac expression domain is indicated by a red arrow. The black arrow indicates the extra sex comb bristles on tarsal segment 2 (TS2). Overexpression of dac in the TS2 segment partially transforms this more distal segment TS2 into the more proximal segment TS1 induces the ectopic sex comb teethes.

Source data

Extended Data Fig. 2 Hi-M library and Pearson correlations of individual replicates.

(a) Schematic representation of the genomic positions of the barcodes of the dac gene locus. Violet bars indicate the positions of PRE1 (barcode 29) and PRE2 (barcode 48). Salmon bar indicates the position of the leg enhancer (RE), green bar indicates the position of the gypsy 2 insertion and gray bar indicates the position of the CG5888 promoter. (b) Scatter plots and Pearson correlations of two biological replicates of wild type pupal leg discs for all tarsal segments and specific tarsal segments (TS1, TS2 and TS3/4). (c) Scatter plots and Pearson correlations of two biological replicates of gypsy 2 mutant pupal leg discs for all tarsal segments and specific tarsal segments (TS1, TS2 and TS3/4).

Extended Data Fig. 3 Insulation activity of gypsy sequences at the dac TAD.

(a) qChIP experiments using Su(HW) antibodies of WT embryos or embryos carrying the gypsy 2 or gypsy 3 insertion. D1-D8, PRE1, 2 and gypsy indicate PCR amplicons used for qChIP experiments along the dac gene locus (see Supplementary Table 2). The housekeeping gene Rp49 was used as negative control. Two independent replicates have been performed. Data are presented as the mean values ± s.d (error bars) of replicates. (b) CUT&RUN profiles for H3K27me3 at the dac gene locus, performed in 3^rd instar imaginal leg discs of the indicated fly lines. Red bar indicates position of the RE. Violet bars indicate positions of PREs. Green bar position of the gypsy insulator. Grey bar below H3K27me3 tracks demarcates the PcG TAD²². (c) The scatter plots show the H3K27me3 enrichment in the 131 Drosophila Polycomb domains in gypsy 1 (left), gypsy 2 (middle) and gypsy 3 (right) as a function of WT flies (log2 scale). The dac Polycomb domain is shown in blue.

Extended Data Fig. 4 Statistical analysis of the insulation changes at the dac domain in the gypsy lines.

(a) The insulation profile per each condition is shown at 3kb resolution as the mean value (line) +/− the standard deviation (shaded area) over the insulation scores computed using 5 different values of the window parameter (w=100, 150, 200, 250, and 300 kb). Each panel show the WT together with one gypsy line at the time: Gypsy1, Gypsy2, and Gypsy3 respectively. These panels also show the locations of the 13 2kb-regions (R0-R12) spaced by 4 kb, spanning the domain from the TAD boundary to the PRE1 (chr2L:16,348,000-16,422,000) on which the differential statistical analysis has been applied. The p-values resulted from a Welch t-test between the WT condition and each of the gypsy mutant at the corresponding locus. (b) Bar-plots of the statistical analysis applied to the R0-R12 regions and the gypsy insertion sites. We notice that the only statistically significant increases of insulation are quantified at the Gypsy 2 and Gypsy 3 insertion sites in the corresponding lines.

Extended Data Fig. 5 Gypsy 1 insertion has insulation activity and reduces PRE looping.

(a) Differential 4C-seq profiles at the dac gene locus of the indicated fly lines using the leg enhancer (RE) as a viewpoint (gray bar). (b) Differential 4C-seq profiles at the dac gene locus of the indicated fly lines using the PRE2 as a viewpoint (gray bar). Reads of 3 replicates were aligned and pooled into 1 kb bins. Mean values of the indicated mutant line are subtracted from the control line and the calculated counts per million are plotted. Green bar indicates position of the gypsy insulator. Position of PREs is indicated by blue dotted lines.

Extended Data Fig. 6 Developmental stage specific consequences of gypsy 2 insertion for the dac Polycomb domain architecture analyzed by 4C-seq.

(a) Differential 4C-seq profiles at the dac gene locus of gypsy 2 versus WT fly lines using the dac promoter (PRE2) as a viewpoint (gray bar) at the indicated developmental stages. (b) Differential 4C-seq profiles at the dac gene locus of gypsy 2 versus WT fly lines using the leg enhancer (RE) as a viewpoint (gray bar) at the indicated developmental stages. Reads of 3 replicates were aligned and pooled into 1 kb bins. Mean values of the indicated mutant line are subtracted from the control line and the calculated counts per million are plotted. Green bar indicates position of the gypsy insulator. Position of PREs is indicated by blue dotted lines.

Extended Data Fig. 7 Consequences of gypsy 2 insertion on chromatin topology observed by Hi-M experiments.

(a) Ensemble Hi-M PWD difference matrix of WT and gypsy 2 mutant for all tarsal segments, TS1, TS2 and TS3/4 of pupal leg discs. Blue indicates shorter distances in WT and red in gypsy2 mutant condition. Green bar indicates the position of the gypsy insulator. Blue square indicates loss of long-range interactions around the position of the PRE1-PRE2 loop upon gypsy insertion. Red dashed square indicates increased short-range interactions up- and downstream the gypsy insulator insertion. (b) Virtual 4C plots derived from Hi-M PWD matrices using barcode 48 (dac TSS) as viewpoint. Green bar indicates the gypsy insulator insertion. Red tracks correspond to WT and blue tracks to gypsy 2 mutant condition.

Extended Data Fig. 8 Cell to cell variability of PRE-PRE or E-P distances measured by Hi-M.

(a) Distances distributions between PRE1 and PRE2, RE and dac TSS and RE and CG5888 TSS. The distances distributions were extracted for TS1 (red), TS2 (orange) and TS3/4 (blue) segments of the WT condition. Asterisk represent the p-value obtained with the Wilcoxon two-sided rank test: p-value [*] < 0.05 (rejected null hypothesis). (b) Distances distributions between PRE1 and PRE2, RE and dac TSS and RE and CG5888 TSS. The distances distributions were extracted for TS1 (red), TS2 (orange) and TS3/4 (blue) segments of the gypsy 2 condition. Asterisk represent the p-value obtained with the Wilcoxon two-sided rank test: p-value [*] < 0.05 (rejected null hypothesis). White box represents the median, the thick black line represents the first and third quartiles, and the red/orange/blue areas represent the kernel density estimation of the distribution.

Extended Data Fig. 9 Importance of the RE for the activation of the CG5888 gene upon loss of the PRE loop.

(a) RT-qPCR analysis at the indicated developmental stages using primers specific for Idgf1, 2 and 3 genes. The fold change between the indicated mutant and wild type (control) line is shown. Three independent experiments have been performed. Data are presented as the mean values ± s.d (error bars) of replicates. * indicates p-value < 0.1, ** indicates p-value < 0.01 (two-sided unpaired t-test). (b) RNA FISH images of WT, gypsy2 and gypsy3 insertion, in 3^rd instar imaginal leg discs (CG5888 gene: green). White bar indicates 30 micrometers. (c) Quantification of CG5888 RNA FISH signals in WT and gypsy2 and gypsy3 imaginal leg discs. The number of CG5888 FISH signals has been quantified and normalized by the surface of the total disc area. Y axis represents the average of 6 imaginal discs values. Error bars indicate s.d. ** indicates p-value < 0.01 (two-sided unpaired t-test).

Source data

Extended Data Fig. 10 Function of PRE loop on gene expression.

(a) A Model illustrating the architectural consequences of gypsy insulator insertion on PRE looping and RE communication with gene promoters. (b) Overlap of Tethering Elements (TEs) identified¹⁵ in and embryonic PREs from⁴⁷.

Supplementary information

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Supplementary Table 1

Coordinates of barcodes used for Hi-M experiments.

Supplementary Table 2

Primers used in this study (qPCR, cloning, CRISPR, RNA FISH and 4C).

Supplementary Table 3

Genomic position of deleted or inserted DNA sequences.

Supplementary Table 4

Hi-C sequencing statistics.

Supplementary Table 5

Number of successfully aligned reads for 4C-seq experiments.

Supplementary Table 6

Differential quantification of H3K27me3 levels within Polycomb domains.

Source data

Source Data Fig. 1

Raw data for graphs.

Source Data Fig. 5

Raw data for graphs.

Source Data Fig. 7

Raw data for graphs.

Source Data Extended Data Fig. 1

Raw data for graphs.

Source Data Extended Data Fig. 9

Raw data for graphs.

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Denaud, S., Bardou, M., Papadopoulos, GL. et al. A PRE loop at the dac locus acts as a topological chromatin structure that restricts and specifies enhancer–promoter communication. Nat Struct Mol Biol (2024). https://doi.org/10.1038/s41594-024-01375-7

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Received: 16 November 2023
Accepted: 22 July 2024
Published: 16 August 2024
DOI: https://doi.org/10.1038/s41594-024-01375-7
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A PRE loop at the dac locus acts as a topological chromatin structure that restricts and specifies enhancer–promoter communication

Abstract

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Main

Results

dac expression during development upon loss of PRE looping

The PRE loop acts as a constitutive 3D chromatin scaffold

A ring enhancer (RE) is necessary and sufficient for dac expression

Loss of dac expression and PRE looping

Gypsy insertion induces insulation and reduces PRE looping

Gypsy insulator does not block RE function

Unchanged enhancer–promoter proximity upon dac gene activation

The PRE loop contributes to enhancer–promoter specificity

Discussion

Gypsy insulator reduces PRE looping but not enhancer function

PRE contacts constitute a specific form of TE loops

PRE looping as a regulator of enhancer–promoter communication

Methods

Fly work and generation of mutant flies by CRISPR–Cas9 genome engineering

Immunostaining experiments

RNA FISH experiments

Hi-C experiments

Hi-C analysis

Hi-M library preparation

Hi-M library hybridization

Hi-M imaging system

Acquisition of Hi-M datasets

Image processing and Hi-M analysis

4C-seq experiments

4C-seq processing and analysis

qRT–PCR experiments

qCHIP experiments

CUT&RUN experiments

CUT&RUN analysis

Reporting summary

Data availability

Code availability

References

Acknowledgements

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Contributions

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Extended data

Supplementary information

Source data

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