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. 2024 Aug 7;13:RP94114. doi: 10.7554/eLife.94114

Stem-loop and circle-loop TADs generated by directional pairing of boundary elements have distinct physical and regulatory properties

Wenfan Ke 1,, Miki Fujioka 2,, Paul Schedl 1,, James B Jaynes 2,
Editors: Yukiko M Yamashita3, Claude Desplan4
PMCID: PMC11305674  PMID: 39110491

Abstract

The chromosomes in multicellular eukaryotes are organized into a series of topologically independent loops called TADs. In flies, TADs are formed by physical interactions between neighboring boundaries. Fly boundaries exhibit distinct partner preferences, and pairing interactions between boundaries are typically orientation-dependent. Pairing can be head-to-tail or head-to-head. The former generates a stem-loop TAD, while the latter gives a circle-loop TAD. The TAD that encompasses the Drosophila even skipped (eve) gene is formed by the head-to-tail pairing of the nhomie and homie boundaries. To explore the relationship between loop topology and the physical and regulatory landscape, we flanked the nhomie boundary region with two attP sites. The attP sites were then used to generate four boundary replacements: λ DNA, nhomie forward (WT orientation), nhomie reverse (opposite of WT orientation), and homie forward (same orientation as WT homie). The nhomie forward replacement restores the WT physical and regulatory landscape: in MicroC experiments, the eve TAD is a ‘volcano’ triangle topped by a plume, and the eve gene and its regulatory elements are sequestered from interactions with neighbors. The λ DNA replacement lacks boundary function: the endpoint of the ‘new’ eve TAD on the nhomie side is ill-defined, and eve stripe enhancers activate a nearby gene, eIF3j. While nhomie reverse and homie forward restore the eve TAD, the topology is a circle-loop, and this changes the local physical and regulatory landscape. In MicroC experiments, the eve TAD interacts with its neighbors, and the plume at the top of the eve triangle peak is converted to a pair of ‘clouds’ of contacts with the next-door TADs. Consistent with the loss of isolation afforded by the stem-loop topology, the eve enhancers weakly activate genes in the neighboring TADs. Conversely, eve function is partially disrupted.

Research organism: D. melanogaster

Introduction

Chromosomes in multicellular animals are organized into a series of topologically independent looped domains, called TADs, or topologically associating domains (Cavalheiro et al., 2021; Chetverina et al., 2017; Jerković et al., 2020; Matthews and White, 2019; Rowley and Corces, 2018). The arrangement of TADs in a given chromosomal DNA segment is generally (though not precisely) similar in different tissues and developmental stages, and this is a reflection of the mechanism underlying TAD formation—the endpoints of TADs are determined by a special class of elements called chromatin boundaries or insulators. While boundary-like elements have been identified in a wide range of animals and plants, the properties of this class of DNA elements have been most fully characterized in Drosophila (Cavalheiro et al., 2021; Chetverina et al., 2017). Fly boundaries have one or more large (100–400 bp) nucleosome-free nuclease-hypersensitive sequences that are targets for multiple DNA binding chromosomal architectural proteins. While only a single chromosomal architectural protein, CTCF, has been characterized in mammals, there are several dozen such proteins in flies, and the list is still growing (Heger et al., 2013; Heger and Wiehe, 2014; Schoborg and Labrador, 2010). In addition to subdividing the chromosome into a series of loops, fly boundary elements have insulating activity. When placed between enhancers or silencers and their target promoters, boundaries block regulatory interactions (Bell et al., 2001; Chetverina et al., 2014; Chetverina et al., 2017). This activity provides a mechanism for delimiting units of independent gene activity: genes located between a pair of compatible boundaries are subject to regulatory interactions with enhancers/silencers present in the same chromosomal interval, while they are insulated from the effects of enhancers/silencers located beyond either boundary in adjacent regulatory neighborhoods. It is currently thought that their ability to organize the chromosome into topologically independent loops is important for their insulating activity (Cai and Shen, 2001; Gohl et al., 2011; Muravyova et al., 2001).

Studies dating back to the 1990s have suggested that fly boundaries subdivide the chromosome into loops by physically pairing with each other (Chetverina et al., 2014; Chetverina et al., 2017). In these first experiments, regulatory interactions were observed for transgenes inserted at distant sites in chromosome that were carrying either the gypsy transposon boundary su(Hw) or the bithorax complex (BX-C) boundary Mcp (Muller et al., 1999; Sigrist and Pirrotta, 1997; Vazquez et al., 1993). Further support for the idea that boundaries function by pairing has come from chromatin immunoprecipitation, chromosome conformation capture (CCC), MicroC, and direct imaging experiments (Chen et al., 2018; Li et al., 2011; Vazquez et al., 2006). More recent studies have revealed physical interactions in the CNS, such as those found for su(Hw) and Mcp, which ‘reach over’ multiple intervening TADs, consistent with them playing an important role in cell type-specific gene regulation (Mohana et al., 2023) by bringing distant enhancers and promoters together.

The parameters governing pairing interactions have been defined using insulator bypass, transvection, and boundary competition assays. These studies have shown that fly boundaries are able to pair not only with heterologous boundaries but also with copies of themselves. Moreover, the pairing interactions typically exhibit a number of characteristic features: promiscuity coupled with clear partner preferences, and orientation dependence.

Partner preferences depend upon the chromosomal architectural proteins that interact with each boundary. For example, in the boundary bypass assay, a set of enhancers are placed upstream of two reporters (Cai and Shen, 2001; Kyrchanova et al., 2008a; Muravyova et al., 2001). When multimerized dCTCF sites are placed between the enhancers and the closest reporters, both reporters are insulated from the enhancers. When a second set of multimerized dCTCF sites are placed downstream of the closest reporter, bypass is observed. In this case the closest reporter, which is bracketed by the multimerized dCTCF sites, is still insulated from the enhancers; however, the downstream reporter is activated (Muravyova et al., 2001). Heterologous combinations give a different result: when multimerized dCTCF sites are placed upstream of the closest reporter and multimerized Zw5 sites are placed downstream, no bypass is observed. Endogenous fly boundaries also show partner preferences in bypass assays and in boundary competition experiments (Gohl et al., 2011; Kyrchanova et al., 2011; Kyrchanova et al., 2008b). On the other hand, while boundaries have partner preferences, they are also promiscuous in their ability to establish functional interactions with other boundaries. For example, the Fab-8 insulator can partner with scs’ from the Drosophila heat shock locus (Gohl et al., 2011).

In addition, to partner preferences, pairing interactions between endogenous fly boundaries are, with a few exceptions, orientation-dependent. Self-pairing interactions are head-to-head. This seems to be a common feature of fly boundaries and has been observed for scs, scs’, iA2, wari, Mcp, Fab-8, AB-I, homie, and nhomie (Fujioka et al., 2016; Kyrchanova et al., 2008a). In contrast, pairing interactions between heterologous boundaries can be head-to-head or head-to-tail. The two boundaries bracketing the even-skipped (eve) locus, homie and nhomie, pair with each other head-to-tail, while boundaries in the Abdominal-B (Abd-B) region of the BX-C usually pair with their neighbors head-to-head. The topology of the loops (TADs) generated by head-to-tail and head-to-head pairing in cis between neighboring boundaries is distinct. As illustrated in Figure 1, head-to-tail pairing generates stem-loops, while head-to-head pairing generates circle-loops. The loops could be connected to each other by unanchored loops (Figure 1A and C), or they could be linked directly to each other if boundaries can pair simultaneously with both neighbors (Figure 1B and D). An alternating pattern of TADs connected by DNA segments that crosslink to each other with reduced frequency (c.f., λ DNA below) is not often observed in MicroC experiments. Instead, most TADs appeared to be directly connected to both of their neighbors without an intervening unanchored loop (Batut et al., 2022; Bing et al., 2024; Levo et al., 2022; see also below). This would suggest that TAD boundaries are typically linked to both neighbors, either simultaneously or as alternating pair-wise interactions.

Figure 1. Diagram of the possible loop topologies generated by head-to-head and head-to-tail pairing.

Figure 1.

(A) Head-to-tail boundary pairing (arrows) generates a series of stem-loops linked together by an unanchored loop. In this case, the main axis of the chromosome would correspond to the unanchored loops connecting different stem-loops. (B) If boundaries pair with both neighbors (head-to-tail), the stem-loops would be linked to each other by the paired boundaries. In this case the main axis of the chromosome would correspond to the paired boundaries. (C) Head-to-head boundary pairing generates a series of circle-loops linked together by an unanchored loop. The unanchored loop will be the main axis of the chromosome. (D) If boundaries pair with both neighbors (head-to-head), the chromatin fiber will be organized into a series of circle-loops connected to each other at their base, and these paired boundaries will define the chromosomal axis. In both (B) and (D), the pairing interactions between the blue and red boundaries need not be in register with the pairing of the red boundary to the next-door green boundary. In this case, the main axis of the chromosome may bend and twist, and this could impact the relative orientation of the stem-loops/circle-loops. More complex structures would be generated by mixtures of stem-loops and circle-loops.

Key to understanding the 3D organization of chromosomes in multicellular eukaryotes will be the identification of TADs that are stem-loops and TADs that are circle-loops. In the studies reported here, we have used MicroC to analyze the contact maps generated by stem-loops and circle-loops. Stem-loop and circle-loop TADs are expected to interact differently with their neighbors, and this should be reflected in the patterns of crosslinking events between neighboring TADs. As illustrated for linked stem-loops in Figure 1B, TAD2 is isolated from its next-door neighbors, TAD1 and TAD3. In this configuration, crosslinking events between sequences in TAD2 and sequences in TAD1 and TAD3 will be suppressed. On the other hand, TAD1 and TAD3 are in comparatively close proximity, and crosslinking between sequences in these two TADs is expected to be enhanced. A different pattern of neighborly interactions is expected for circle-loop TADs. In this case, the TAD in the middle, TAD2, is expected to interact with both of its neighbors (Figure 1D). To test these predictions, we have first compared the MicroC contact profiles for stem-loop and circle-loop TADs. For stem-loops we selected the eve TAD, while for circle-loops we chose the four TADs that comprise the Abd-B parasegment-specific regulatory domains. We show that these stem-loop and circle-loop TADs have distinctive crosslinking signatures. To confirm these MicroC signatures, we converted the topology of the eve TAD from a stem-loop to a circle-loop. In addition to changing the MicroC signature of the eve TAD, the change in topology is accompanied by changes in the regulatory interactions between eve and its neighbors.

Results

Stem-loops versus circle-loops

The distinctive loop topologies of stem-loops and circle-loops are expected to be reflected in the contact maps that are generated in MicroC experiments. To determine if this is the case, we compared the MicroC contact maps for the eve TAD and the TADs that correspond to the four Abd-B parasegment-specific regulatory domains, iab-5, iab-6, iab-7, and iab-8. The eve TAD is generated by pairing interactions between the nhomie boundary upstream of the eve transcription unit and the homie boundary downstream. Since nhomie and homie pair with each other head-to-tail, the eve TAD has a stem-loop topology (Fujioka et al., 2016). Unlike the eve boundaries, the boundaries that delimit the Abd-B regulatory domains are thought to pair with their neighbors head-to-head (Chetverina et al., 2017; Kyrchanova et al., 2008a; Kyrchanova et al., 2011; Kyrchanova et al., 2008b). This means that the parasegment-specific regulatory domain TADs, iab-5, iab-6, iab-7, and iab-8, are expected to have a circle-loop topology.

As shown in Figure 2, the eve TAD and the four Abd-B TADs have distinctive MicroC contact patterns. The eve TAD is a ‘volcano’ triangle with a plume. The endpoints of the volcano triangle are delimited by nhomie on the left and homie on the right, and within the eve locus (the volcano), there are additional enhanced interactions. While the volcano triangle is generated by contacts between sequences within the eve stem-loop, contacts between sequences in eve and in the neighboring TAD on the left, TL (which contains multiple sub-TADs and six genes: CG15863, CG1418, Pal1, CG12133, eIF3j, and CG12134), are much reduced (L-ev in Figure 2A). There is a similar suppression of contacts between sequences in the eve TAD and sequences in the large neighboring TAD on the right, TM (which contains TER94 and Pka-R2; ev-M in Figure 2A). On the other hand, as expected from the regulatory interactions observed for stem-loops in boundary bypass experiments (Kyrchanova et al., 2008a), physical contacts between sequences in TL and TM are enhanced compared to those between eve and TL (L-ev) or TM (ev-M). Because of the preferential interactions between TADs to either side of the eve stem-loop, the eve volcano triangle is topped by a plume (L-M in Figure 2A). TM also interacts with the two TADs farther to the left of eve, TK (K-M in Figure 2A), and TJ (J-M in Figure 2A; see also Figure 2—figure supplement 1).

Figure 2. Stem-loops and circle-loops.

Once in a while you get shown the light in the strangest of places if you look at it right.”a MicroC contact profile for Drosophila wild-type (yw) NC14 embryos. The bin size for each panel is 200 bp. (A) eve and neighboring TADs (TI, TJ, TK TL, TM, and TN). The eve TAD is a volcano with a plume that is anchored by nhomie (nh) and homie (h). The plume is generated by crosslinking of sequences in the two neighboring TADs. At the bottom of the plume, TL sequences are linked to sequences in TM close to eve, including TER94. At the next level, sequences in TK are linked to TM (region K-M). In addition, sequences in TL are linked to sequences in TM located beyond the TER94 gene. At the next level, sequences in TJ are linked to sequences in TM. Note that interactions between sequences in TL and TJ and sequences in TM close to the eve TAD are somewhat less frequent than those farther away from the eve TAD. Sequences in the neighboring TADs also interact with each other, as indicated. For example, sequences in TK and TJ interact with each other (J–K) and also interact with sequences in TI (I–K and I–J). (B) The BX-C gene Abd-B and the parasegment- (PS-) specific regulatory domains iab-3, iab-4, iab-5, iab-6, iab-7, and iab-8. iab-4 regulates the abd-A gene in PS9, while iab-5 – iab-8 regulate Abd-B in PSs 10–13, respectively. These domains are separated from each other by the boundary elements Fab-4, Mcp, Fab-6, Fab-7, and Fab-8, as indicated. The AB-I boundary is located upstream of the Abd-B promoter. Each regulatory domain corresponds to a TAD. Though partially insulated from each other, each TAD interacts with its immediate neighbors. For example, iab-5 interacts with its immediate neighbors iab-4 and iab-6 to give 4–5 and 5–6, respectively. It also interacts with the next-next-door neighbor iab-7 (5-7) and even its next-next-next-door neighbor iab-8 (5-8). (a From ‘Scarlet Begonias’ by the Grateful Dead, 1974).

Figure 2.

Figure 2—figure supplement 1. MicroC contact profiles for nhomie forward, lambda DNA, nhomie reverse, and homie forward in larger scale.

Figure 2—figure supplement 1.

N (replicates) = 2. Resolution = 200. (A) MicroC contact maps for the nhomie forward replacement. (B) MicroC contact maps for the lambda DNA replacement. (C) MicroC profile of the nhomie reverse replacement. (D) MicroC profile of the homie forward replacement.

Like the eve boundaries, the TADs in the Abd-B region of BX-C region are connected to their neighbors by the boundaries at their base. As predicted from genetic studies on BX-C boundaries, each TAD corresponds to one of the four parasegment-specific regulatory domains. While the MicroC contact maps for the four Abd-B TADs resemble the contact patterns in the eve TAD, these Abd-B TADs differ from eve in that there are no plumes above their triangle peaks (Figure 2B). Instead, the Abd-B TADs are overlaid by a series of rectangular interlocking low-density contact (LDC) domains, or clouds. As illustrated in Figure 2B, the iab-6 regulatory domain is flanked by clouds generated by crosslinking with its next-door neighbors iab-5 (5–6) and iab-7 (6–7), followed by crosslinking with neighbors that are a TAD away from iab-6, iab-4 (4–6), and iab-8 (6–8). The other regulatory domains also form a unique set of interlocking LDCs/clouds with their immediate neighbors, their next-next-door neighbors, and their next-next-next-door neighbors.

TAD formation in a nhomie deletion

To further investigate the pairing properties and functioning of the eve boundaries, we used CRISPR-Cas9 to add two attP sites flanking the nhomie region, replacing the region with a mini-white gene. Using mini-white as an exchange marker, ΦC31 recombinase-mediated cassette exchange (RMCE) was used to restore the sequence of the region, with nhomie modifications. As a control for possible effects of the sequences introduced in generating the modification, we reinserted a 597 bp nhomie DNA fragment in the same orientation as the endogenous nhomie boundary (nhomie forward). To maintain roughly the same distance between eve and the neighboring TAD in the nhomie deletion, we introduced a 606 bp DNA fragment from phage λ (λ DNA). Figure 3 (and Figure 2—figure supplement 1A and B) shows the MicroC contact profiles for the nhomie forward and λ DNA replacements in 12–16 hr embryos (mid-embryogenesis: stages 12–14). Except that the sequencing depth of the nhomie forward replacement is not as great as the WT shown for the eve locus in Figure 2, the profile is quite similar. Like WT, there are sub-TADs within the eve TAD. One of these appears to link homie and the neighboring PRE to the eve promoter-proximal PRE (Fujioka et al., 2008), and is marked by an interaction dot (asterisk in Figure 3A). Another links nhomie to the eve promoter region (blue arrow in Figure 3A). The eve TAD is topped by a plume, which is generated by interactions between sequences in neighboring TADs TL with TM (L-M). On the other hand, interactions between eve and its neighbors are suppressed. Like eve, there are sub-TADs in the neighboring TAD, TL. The TL sub-TAD closest to eve (TL4) corresponds to the CG12134 transcription unit (green arrowhead marks the boundary: Figure 3), while the neighboring sub-TAD (TL3) encompasses the eIF3j transcription unit (blue arrowhead: Figure 3).

Figure 3. TAD organization of the nhomie forward and lambda DNA replacements.

Figure 3.

(A) MicroC contact profile of 12–16 hr embryos (stage 12–14) nhomie forward embryos. In this, our positive control, nhomie replaces endogenous nhomie, in the same orientation. N (replicates) = 2. Resolution = 200 bp. L-M: interactions between TADs TL and TM flanking the eve locus. Asterisk: sub-TAD linking the eve promoter to the eve PRE and homie. Dark blue arrow: sub-TAD linking the eve promoter to nhomie. Light blue arrow: nhomie. Red arrow: homie. Green arrowhead: sub-TAD boundary formed by the CG12134 promoter region. Dark blue arrowhead: sub-TAD boundary formed by eIF3j promoter region. Diagram: map of eve locus and surrounding genes. (B) Virtual 4C with viewpoint from homie (black arrow) in nhomie forward embryos. (C) Diagram of the eve stem-loop TAD. (D) MicroC contact profile of 12–16 hr λ DNA embryos. In this replacement, λ DNA is inserted in place of nhomie. N (replicates) = 3. Resolution = 200 bp. Asterisk: sub-TAD linking the eve promoter to the eve PRE and homie. Purple arrowhead: sub-TAD linking CG12134 promoter region to the eve promoter. The eIF3j sub-TAD TL3 (between the blue and green arrowheads) is still present. (E) Virtual 4C with viewpoint from homie (black arrow) in λ DNA embryos. (F) Diagram of the ‘unanchored’ eve TAD. Double arrows show novel interactions.

The MicroC profile of the λ DNA replacement (Figure 3D) is quite different from that of either nhomie forward or WT, which are similar (Figures 2A and 3A). While homie still defines the distal (relative to the centromere) end of the eve locus, the λ DNA replacement does not function as a TAD boundary, and the leftward endpoint of the eve TAD is no longer well-defined. One new ‘endpoint’ for the eve locus maps to sequences between CG12134 and eIF3j (green arrowhead), which in wild type corresponds to the left boundary of the TL sub-TAD TL4. The other endpoint maps to sequences between eIF3j and CG12133 (blue arrowhead), which in wild type define the left boundary of the TL sub-TAD TL3. These interactions are not as stable as those between nhomie and homie as the density of interaction dots is lower. Furthermore, they appear to flip back and forth between alternative endpoints (as indicated by the green and blue double arrows in Figure 3F) based on the MicroC contact profile, which is consistent with a mixture of (at least) two conformations. The eve TAD also interacts with sequences in the two other TL sub-TADs, TL1 and TL2. In addition, the eve promoter appears to interact with sequences located upstream of CG12134 (purple arrowhead in Figure 3D and double arrow in Figure 3F), while this interaction is not observed in the nhomie forward replacement. While the TL TAD (from the TK:TL boundary to nhomie) is also disrupted by the nhomie deletion (it has a much less distinct ‘volcano apex’, and its right-most sub-TAD TL4 is now fused with the eve TAD), the left-most TL sub-TADs (TL1, TL2, and TL3) are still present, indicating that their formation does not depend on nhomie. As shown in the virtual 4C at homie viewpoint in Figure 3B and E, the homie boundary interacts with the nhomie forward replacement, but does not contact the λ DNA replacement.

eve enhancers activate eIF3j expression in the nhomie deletion

In transgene assays, boundary elements block regulatory interactions when interposed been enhancers (or silencers) and reporter genes (Chetverina et al., 2014; Chetverina et al., 2017; Kellum and Schedl, 1992). To determine whether this is also true in their endogenous context, we compared the expression in syncytial blastoderm embryos of the two genes that flank nhomie at the eve locus, eIF3j and CG12134. As the nhomie deletion eliminates homie’s pairing partner and disrupts the eve TAD, we also examined the expression of eve and of the gene just beyond the homie boundary, TER94, which has strong maternal expression through stage 11 (Figure 4—figure supplement 2B, WT). Consistent with the seemingly normal MicroC profile on the homie side of the eve TAD, we did not detect evidence of eve-like TER94 expression (Figures 4 and 5). Thus, the formation and functioning of the TM TAD do not appear to be impacted by either the loss of nhomie per se or the fact that the left end of the eve TAD is no longer properly anchored.

Figure 4. nhomie deletion (λ DNA replacement) exposes eIF3j to the eve enhancers.

nh forward: positive control, as in Figure 3. λ DNA: nhomie is replaced with λ DNA. At the syncytial blastoderm stage, a series of stripe-specific enhancers upstream (stripes 1, 2, 3, 7) and downstream (stripes 1, 4, 5, 6) of the eve gene drive eve expression. During cellularization of the blastoderm and gastrulation, a single enhancer located upstream of eve drives expression of all seven stripes. DAPI: DNA stained with DAPI (blue). eIF3j: embryo hybridized with probe complementary to eIF3j mRNA. eve: embryo hybridized with probe complementary to eve mRNA. TER94: embryo hybridized with probe complementary to TER94. Yellow arrowheads: eve-enhancer-driven eIF3j stripes. Control nonspecific probes for each channel indicate autofluorescence background in the top panel. Scale bar = 100 µm.

Figure 4.

Figure 4—figure supplement 1. Expression of CG12134 in WT (yw) and the four nhomie replacements.

Figure 4—figure supplement 1.

Digoxigenin in situ hybridization was used to detect expression of CG12134 during development in the indicated genetic backgrounds. Approximate developmental stages of the embryos in each genetic background are shown on the right. As controls, embryos of similar stages were hybridized with an eve probe. Scale bar = 50 µm.
Figure 4—figure supplement 2. Expression of eIF3j (Adam) and TER94 in WT (yw) and the four nhomie replacements.

Figure 4—figure supplement 2.

Digoxigenin in situ hybridization was used to detect expression of (A) eIF3j and (B) TER94 during development in the indicated genetic backgrounds. Approximate developmental stages of the embryos in each genetic background are shown on the right. As controls, embryos of a similar stage were hybridized with an eve probe. Note that unlike the HCR-FISH results shown in Figure 4, we can detect a low level of eIF3j stripe expression in WT (yw) and nhomie forward stage 5 embryos. The difference is likely due to the fact that signal amplification in the digoxigenin in situ hybridization procedure is nonlinear, resulting in greater contrast between background and signal when staining conditions are optimal. Scale bars = 50 µm.

Figure 5. Manipulating the nhomie boundary impacts the regulatory landscape.

Figure 5.

N = # of independent replicates, n = # of embryos. Two-way ANOVA with Tukey’s multiple comparisons test for each pair of groups was used to determine the statistical significance. *p≤0.05, **p≤0.01, ***p≤0.001, and ****p≤0.0001. (A) Quantitation of the number of embryos showing stripe patterns in HCR-FISH for eIF3j and TER94, as shown in Figures 4 and 7. N = 3. n = 45 for each group. (B) Quantitation of the number of missing ventral denticle bands in larvae from a cross of BSC/CyO,hb-lacZ deficiency females to males of the indicated genotypes (N = 6): wild-type control (yw), n = 767. For the nhomie forward replacement, n = 1099; for the λ DNA replacement, n = 1175; for the nhomie reverse replacement, n = 1083; for the homie forward replacement, n = 1137.

In the case of the gene closest to the nhomie deletion, CG12134, we were unable to consistently detect transcription driven by the eve enhancers in either nhomie forward or λ DNA embryos. In some λ DNA embryos, there were hints of stripes at the blastoderm stage (see Figure 4—figure supplement 1); however, these ‘stripes’ were not observed in most embryos. Since CG12134 (which forms the TL4 sub-TAD in wild type) is closest to the eve enhancers and interacts most strongly, it is possible that the promoter is not compatible with the eve enhancers. A different result was obtained for eIF3j in λ DNA embryos. As shown in the HCR-FISH experiment in Figure 4 and quantitated in Figure 5, we observed a series of eIF3j stripes over a dark background in pre-cellular blastoderm embryos. As this background hybridization is evident in earlier stages, much of it is likely to be of maternal origin. In contrast to the λ DNA replacement, these stripes are not visible in the nhomie forward (control) replacement (Figures 4 and 5A). While it is possible to detect all seven stripes in λ DNA blastoderm stage embryos, their levels of expression are not equal. The highest levels correspond to eve stripes 1, 2, 3, and 7, while eve stripes 4, 5, and 6 are expressed at much lower levels. Since the stripe enhancers for 1, 2, 3, and 7 are located between the eve promoter and nhomie, they are closer to the eIF3j promoter than the enhancers for stripes 4, 5, and 6, which are located downstream of the eve transcription unit. In addition to possible effects of distance, the subdomain linking the eve PRE and homie to the promoter is still observed in the λ DNA replacement, and this could partially sequester the enhancers located downstream of the eve promoter.

We also used digoxigenin in situ hybridization to analyze eIF3j expression. With this procedure we were able to detect a low level of eve-activated eIF3j stripe expression in WT and nhomie forward embryos at stage 5 (blastoderm) and stages 7–8 (early gastrula: Figure 4—figure supplement 2A). In stage 5 embryos when eve expression is driven by specific stripe enhancers, eIF3j expression appears to be similar in all seven stripes (Figure 4—figure supplement 2, WT and nhomie forward). In contrast, as was observed in the HCR-FISH experiments (Figure 4), there is a clear bias for enhancers located upstream of eve, where eIF3j is also located, in the λ DNA replacement at this point in development. In stage 7–8 embryos, the seven-stripe enhancer drives eve expression. It is located close to nhomie and, not surprisingly, high levels of eIF3j expression are observed in all seven stripes in the λ DNA replacement (Figure 4—figure supplement 2A). At later embryonic stages, eve expression is driven by tissue-specific neurogenic, mesodermal, and anal plate enhancers. However, eIF3j is expressed at high levels in a complex pattern in older embryos, and we were unable to unambiguously detect expression driven by the eve enhancers over this background mRNA. This is also not surprising, given that all of the enhancers for these aspects of eve expression are located downstream of the eve promoter, like the enhancers driving stage 5 stripes 4, 5, and 6.

While the nhomie deletion did not have any obvious impact on the level or pattern of eve expression in blastoderm stage embryos (see Figure 4), it seemed possible that eve activity was not entirely normal. To test this possibility, we mated males homozygous for either λ DNA or nhomie forward to females heterozygous for a chromosomal deficiency that includes the eve gene. We then quantitated the number of missing denticle bands in embryonic cuticle preps. As shown in Figure 5B, the frequency of larvae with ‘severe’ defects (two or more missing ventral denticle bands) in nhomie forward is similar to that in a WT yw control. In contrast, in the λ DNA replacement, the frequency of larval cuticles with two or more missing denticle bands is increased nearly twofold. Taken together, the increase in severity of the cuticle defects is significant at the p<0.01 level (one-tailed t-test). The A6 denticle band is missing most frequently, followed by A2, A4, and then A8. These even-numbered abdominal denticle bands are those that are lost in eve deficiency mutants (from which the name even skipped comes), suggesting that eve stripe expression at blastoderm stages is compromised in the embryos that produce these defective cuticles.

The eve TAD is converted from a stem-loop to a circle-loop by inverting nhomie

The orientation of boundary:boundary pairing interactions determines the topology of each chromatin loop (Bing et al., 2024; Fujioka et al., 2016). Since nhomie and homie pair with each other head-to-tail, the endogenous eve TAD is a stem-loop. This orientation dependence means that one can convert the eve TAD from a stem-loop to a circle-loop by inverting the nhomie boundary. If our expectations are correct, the MicroC contact pattern will also be transformed from a volcano triangle with a plume to one in which sequences in the eve TAD are flanked by a cloud of crosslinked sequences from both neighboring TADs (TL and TM), like that observed in the Abd-B region of the BX-C.

We tested this prediction by inserting the nhomie boundary in the reverse orientation (nhomie reverse). Figure 6A shows that the eve TAD is reconstituted by nhomie reverse (compare with Figure 3: see also Figure 2—figure supplement 1). The sub-TAD evident in the nhomie forward replacement linking nhomie to the eve promoter is also re-established (blue arrow). In addition, consistent with our expectation, the plume topping the eve TAD is gone and is replaced by a much more sparsely populated LDC domain (purple double-arrow and above). The more prominent LDC TAD-TAD interactions (the clouds) are between sequences in the eve TAD and the neighboring TADs. On the right, eve forms an LDC interaction domain with TM (ev-M). On the left, eve interacts most strongly with sequences in TL4, and progressively less strongly with sequences in the sub-TADs TL3, TL2, and then TL1 (L-ev). In addition to restoring the eve TAD, the TL TAD is re-established, indicating that the nhomie boundary is important in defining both endpoints of the TL TAD. On the other hand, with the exception of the CG12133 sub-TAD, nhomie does not play a role in generating the three other sub-TADs in the TL TAD. The other interesting feature is a 45o band of interaction (just below the purple double-arrow) that includes interactions between sequences in TL4 and sequences in eve that appear to be located near the left edge of homie. These sequences likely correspond to the eve 3′ PRE (Fujioka et al., 2008), located just inside the 3′ end of the eve TAD.

Figure 6. TAD organization of the nhomie reverse and homie forward replacements.

Figure 6.

(A) MicroC contact profile of 12–16 hr nhomie reverse embryos. In this replacement, nhomie is inserted in the reverse orientation compared to WT nhomie. N (replicates) = 3. Resolution = 200 bp. (B) Virtual 4C with viewpoint from homie (black arrow) in nhomie reverse embryos. (C) Diagram of the nhomie reverse:homie circle-loop. (D) MicroC contact profile of 12–16 hr homie forward embryos. In this replacement, homie is inserted in the forward orientation (the same as the endogenous homie): N (replicates) = 3, resolution = 200 bp. (E) Virtual 4C with viewpoint from homie (black arrow) in homie forward embryos. (F) Diagram of the homie forward:homie circle-loop. (A, C) Note that interactions between the TADs flanking the eve locus (purple double arrow) are suppressed compared to nhomie forward (see Figure 3), while interactions of the eve TAD (Tev) with TL and TM are enhanced (L-ev and ev-M). Asterisk: sub-TAD linking the eve promoter to the eve PRE and homie. Dark blue arrow: sub-TAD linking the eve promoter to nhomie reverse. Light blue arrow: nhomie reverse. Red arrow: homie. Green arrowhead: sub-TAD boundary formed by the CG12134 promoter region. Dark blue arrowhead: sub-TAD boundary formed by the eIF3j promoter region.

Insulation is reduced in nhomie reverse

Consistent with the models for stem-loops and circle-loops in Figure 1, the neighborly interactions evident in the MicroC contact patterns for nhomie forward and nhomie reverse are quite distinct. The nhomie forward TAD is isolated from its neighbors, and crosslinking between eve and the neighboring TADs is suppressed (Figures 1B and 2A). This is not true for the circle-loop TAD generated by nhomie reverse: in this configuration, the eve TAD is not sequestered from neighboring TADs (Figures 1D and 2B), but instead interacts much more frequently with sequences in next-door TADs than in the stem-loop configuration. Since the eve TAD is no longer as well-isolated from its neighbors, this could increase the frequency of ‘productive’ interactions between eve regulatory elements and genes in nearby TADs, and vice versa.

To test whether the circle-loop topology has an impact on the regulatory landscape, we hybridized nhomie reverse embryos with HCR-FISH probes for eIF3j, TER94, and eve. In early blastoderm stage nhomie forward embryos, there is little evidence of eIF3j or TER94 expression driven by eve stripe enhancers, and the HCR-FISH hybridization pattern is uniform (see Figure 4). In contrast, it is possible to discern individual stripes of both eIF3j and TER94 mRNA over the background signal in a subset of nhomie reverse blastoderm stage embryos in HCR-FISH (Figure 7). Since these genes are assembled into their own topologically independent looped domains rather than being in the same domain as eve, the level of eIF3j and also TER94 stripe expression is lower than that observed in the nhomie deletion (λ DNA). The nhomie reverse circle-loop also differs from the nhomie deletion (λ DNA) in that there is not such an obvious preference for which eve enhancers activate expression. In addition, eve-dependent eIF3j and TER94 stripes are detected in only about half of the blastoderm stage embryos (Figure 5A). It is possible that the frequency of productive inter-TAD contacts differs from one embryo to the next; however, a more likely reason is that the high background of eIF3j and TER94 transcripts obscures the low level of eve enhancer-driven expression at this stage. Once the seven-stripe enhancer is activated, eve-dependent TER94 expression in nhomie reverse is elevated, and all seven stripes are observed (Figure 4—figure supplement 2B). This fits with the MicroC contact profile. As shown in Figure 6A, the TER94 gene is preferentially crosslinked to eve sequences located between nhomie reverse and the eve promoter compared to sequences spanning the eve gene and the downstream enhancers (i.e., the upper-left portion of the ev-M region shows more crosslinking than does the lower-right portion). This bias correlates with the location of the seven-stripe enhancer, located near the 5′ end of the eve TAD. By contrast, there is much less seven-stripe enhancer-driven expression of eIF3j (Figure 4—figure supplement 2A) than of TER94. Consistent with this observation, crosslinking between eIF3j and sequences in eve close to nhomie reverse and the seven-stripe enhancer occur less frequently than crosslinking to sequences on the other side of the eve TAD (i.e., the lower-left portion of the L-ev region of the MicroC profile shows less crosslinking than does the upper-right portion), although this crosslinking bias may not be as pronounced as that observed between TER94 and the two sides of the eve TAD.

Figure 7. eve enhancers activate neighboring genes when the eve TAD is a circle-loop.

Figure 7.

HCR-FISH hybridization to mRNA expressed by eIF3j, eve and TER94 at the blastoderm stage (embryonic stage 5). nh forward: nhomie is replaced with nhomie in the forward (normal) orientation (positive control, as in Figure 3). nh reverse: nhomie is replaced with nhomie in the reverse orientation. h forward: nhomie is replaced with homie in the forward orientation. Yellow arrowheads: positions of stripes. DAPI (blue): DNA stained with DAPI. eIF3j (green): embryo is hybridized with probe complementary to eIF3j mRNA. eve (orange): embryo is hybridized with probe complementary to eve mRNA. TER94 (red): embryo is hybridized with probe complementary to TER94. Control nonspecific probes for each channel indicate autofluorescence background in the top panel. Scale bar = 100 µm.

While eve stripe expression is not discernibly different from wild type, the circle-loop topology still impacts eve function. As shown in Figure 5B, the fraction of nhomie reverse embryos with two or more missing denticle bands is nearly twice that in either yw or the nhomie forward replacement. Taken together, the increase in severity of the cuticle defects is significant at the p<0.05 level (one-tailed t-test). As was observed for the λ DNA replacement, the A6 denticle band is missing most frequently, followed by A2, A4, and then A8.

homie forward converts the eve stem-loop into a circle-loop

While the findings in the previous section show that loop topology impacts how sequences in TADs interact with each other and with their neighbors, one might argue that the effects we observed are a reflection of some novel properties of the nhomie boundary when it is inverted. To test this possibility, we took advantage of the fact that in addition to pairing with nhomie, the homie boundary pairs with itself (Fujioka et al., 2016). However, unlike nhomie:homie pairing, which is head-to-tail, homie:homie pairing is head-to-head. This means that it is possible to convert the eve TAD into a circle-loop by inserting homie into the nhomie deletion in the forward orientation.

As shown in Figure 6D (Figure 2—figure supplement 1), the MicroC contact profile of homie forward is similar to that observed for nhomie reverse. The plume topping the eve TAD in wild type (Figure 2A) or in nhomie forward (Figure 3A) is absent. Likewise, instead of being isolated from its neighbors, the eve TAD contacts TL and TM. Also like nhomie reverse, homie forward forms a subdomain within the eve TAD linking it to sequences in the eve promoter. There is also enhanced crosslinking between sequences in TL4 and sequences on the right end of the eve TAD that correspond to the eve 3′ PRE (Fujioka et al., 2016).

The MicroC pattern is not the only similarity between homie forward and nhomie reverse. The functional properties of the homie forward eve TAD are also similar. Figure 7 shows that the eve enhancers weakly activate both eIF3j and TER94. As was the case for nhomie reverse, expression levels at the blastoderm stage are low and are observed in only about half of the embryos (Figure 5A). After the blastoderm stage, when the seven-stripe enhancer drives eve expression, an even higher level of TER94 expression is observed (Figure 4—figure supplement 2B). In addition, the functioning of the eve gene when it is in the circle-loop configuration is not as efficient, and the frequency of homie forward embryos with two or more missing denticle bands is twice that of nhomie forward (Figure 5B). Taken together, the increase in severity of the cuticle defects is significant at the p<0.05 level (one-tailed t-test).

Discussion

Two different though overlapping classes of chromosomal architectural elements have been identified in flies. One class is the PREs found in many developmental loci. PREs were first discovered because they induce pairing-sensitive silencing of reporter genes (Americo et al., 2002; Kassis et al., 1991). More recent studies have shown that the ability of these elements to physically pair with each other may be their most important function (Batut et al., 2022; Levo et al., 2022). The other class of architectural elements are chromatin boundaries (also called insulators). PRE pairing in cis typically takes place within the context of a larger chromosomal domain, or TAD. In contrast, boundary elements are responsible for defining the endpoints of these looped domains (Arzate-Mejía et al., 2020; Batut et al., 2022; Bing et al., 2024; Chetverina et al., 2017; Ibragimov et al., 2023; Stadler et al., 2017). While not much is known about the parameters governing PRE pairing, the pairing interactions of fly boundaries have been studied in some detail. The key features include an ability to engage in promiscuous pairing interactions, distinct partner preferences, and orientation dependence. Of the endogenous (non-gypsy) boundaries whose functional properties have been studied in detail, only one, Fab-7, appears to be able to pair in both orientations. However, Fab-7 may be unusual in that its boundary activity depends upon factors that have been implicated in the functioning of PREs (Kyrchanova et al., 2018). For all of the other boundaries studied so far, pairing interactions are orientation-dependent. When fly boundaries pair with themselves, the interactions are head-to-head (Kyrchanova et al., 2008a). This make sense, as the available evidence suggests that self-pairing interactions in trans may be largely responsible for the pairing of homologous chromosomes in precise register (Erokhin et al., 2021; Fujioka et al., 2016). In this case, head-to-tail self-pairing would uncouple the loops on the two homologs (and sister chromatids).

Unlike self-pairing in trans, pairing interactions between heterologous boundaries in cis can be head-to-head or head-to-tail. The topological consequences are quite distinct. The former generates a circle-loop, while the latter forms a stem-loop (Chetverina et al., 2017). In the studies reported here, we have investigated how these two different topologies impact the local chromatin organization. We have also determined whether circle-loops and stem-loops alter the ability of boundary elements to define units of independent gene activity and insulate against regulatory interactions between neighboring TADs.

nhomie deletion disrupts the eve TAD

As would be predicted from many different studies (Cavalheiro et al., 2021; Chetverina et al., 2017), deletion of the nhomie boundary and replacement with a control λ DNA disrupts the eve TAD and alters the regulatory landscape. The MicroC profile shows that disruption of the eve TAD and the neighboring TADs is one-sided. Within the eve TAD, the subdomain linking homie and the nearby PRE to the eve promoter is unaffected. Likewise, the large TAD, TM, which encompassess both TER94 and pka-R2, and is defined at one end by homie and at the other by an uncharacterized boundary element upstream of the pka-R2 promoter, is intact (Figure 2—figure supplement 1B). In contrast, on the nhomie side of the eve TAD, the sub-TAD linking nhomie to the eve promoter is absent and is replaced by a less well-defined sub-TAD linking the eve promoter to an element near the CG12134 promoter. However, the endpoint of the eve TAD is no longer distinct, and eve sequences are now crosslinked to the eIF3j sub-TAD and to sequences in the more distant TL sub-TADs TL3, TL2, and TL1. Consistent with these alterations in the physical organization of the eve and neighboring TADs, the TER94 gene is still insulated from the eve enhancers. While sequences in the eve TAD physically interact with the gene closest to nhomie, CG12134, only the next gene over, eIF3j, is clearly activated by the eve enhancers. Since crosslinking between sequences in CG12134 and the eve TAD is more frequent than crosslinking between the eIF3j sub-TAD TL3 and the eve TAD, it seems likely that CG12134 is refractory to activation by the eve enhancers. This could be due to an incompatibility between the CG12134 promoter and the eve enhancers. Alternatively, the promoter may not be active at this stage. While eIF3j is activated by the eve enhancers in stage 5 embryos in the nhomie deletion, the stripe enhancers located upstream of the eve gene drive a higher level of expression than those located downstream. Two factors in addition to the effects of distance could potentially account for this finding. Since the eve promoter is located between the downstream enhancers and the eIF3j gene, activation of eIF3j by the downstream enhancers could be suppressed by promoter competition. Alternatively, or in addition, the sub-TAD formed between the 3′ PRE/homie and the eve promoter/proximal PRE (Fujioka et al., 2008) could tend to isolate the downstream eve stripe enhancers from interactions with eIF3j.

Topology impacts local 3D genome organization and the potential for regulatory interactions

In boundary bypass experiments using endogenous fly boundaries, the ability of the upstream enhancers to activate the downstream reporter depended on the topology of the loop generated by the paired boundaries (Kyrchanova et al., 2008a). Activation is observed for stem-loops, as this configuration brings the upstream enhancers into close proximity with the downstream reporter. In contrast, the enhancers and downstream reporter are not brought into contact when the topology is a circle-loop. As would be predicted from these bypass experiments, the stem-loop formed by the head-to-tail pairing of nhomie and homie physically isolates the eve TAD from its neighbors, and this is reflected in the low density of contacts between sequences in eve and the neighboring TADs (Figure 2A). Conversely, the TADs that flank eve are brought together, and contacts between them generate the plume that is observed above the eve volcano triangle.

The physical isolation afforded by the stem-loop topology is lost when the eve TAD is converted to a circle-loop either by inverting the nhomie boundary or by replacing nhomie with the homie boundary inserted in the forward direction (Figure 6). In the former case, head-to-tail nhomie:homie pairing generates a circle-loop. In the latter case, head-to-head pairing of homie (inserted in the forward orientation) with endogenous homie generates a circle-loop. The alteration in the local 3D organization induced by the conversion of eve to a circle-loop is evident from the changes in the MicroC contact pattern. Instead of being isolated from neighboring TADs, the eve TAD interacts not only with its immediate neighbors, but also with more distant TADs. As a result, the plume of enhanced contacts linking TM to TL, TK, and TJ is absent and is replaced by contacts between these TADs and the eve TAD.

As might be expected from the MicroC contact patterns, the conversion to a circle-loop topology is accompanied by alterations in regulatory interactions between eve and the genes in the neighboring TADs (Figure 7, Figure 4—figure supplement 2). Unlike the nhomie forward replacement, the eve stripe enhancers in both of the circle-loop replacements are able to weakly activate expression of two neighboring genes, eIF3j and TER94. This pattern of activation mirrors the enhancement in contacts between the eve TAD and the neighboring TL and TM TADs evident in MicroC experiments. Thus, though eIF3j and TER94 are clearly shielded from the eve enhancers when the eve TAD is a circle-loop, a greater degree of isolation from the action of the eve enhancers is afforded when the eve TAD is a stem-loop.

In addition to reducing insulation from regulatory interactions with genes in neighboring TADs, the circle-loop topology impacts the functioning of the eve gene (Figure 5B). For both nhomie reverse and homie forward, the frequency of multiple denticle band defects compared to the nhomie forward control is enhanced in a sensitized genetic background. While we did not detect any obvious reductions in the eve stripes in blastoderm stage embryos, the circle-loop provides less insulation than the stem-loop, and it is possible that the neighboring promoters suppress eve expression by competing for the eve enhancers. Another (nonmutually exclusive) possibility comes from the studies of Yokoshi et al., 2020, who used live imaging to examine the effects of flanking a reporter with the nhomie and homie boundaries. In their experiments, reporter expression was enhanced over twofold when the reporter was flanked by nhomie and homie; however, the enhancement was greater when the paired boundaries formed a stem-loop than when they formed a circle-loop.

Boundary:boundary pairing can generate stem-loops and circle-loops

Our manipulations of the nhomie boundary support the notion that TADs can have two different loop topologies, stem-loop and circle-loop, and these topologies impact their physical and biochemical properties. Since circle-loop TADs cannot be generated in the popular cohesin loop extrusion/CTCF roadblock model for the sculpting the 3D genome, it would be important to know whether there are other unambiguous examples of loops with either a stem-loop or circle-loop topology besides those described here. As discussed above, the available evidence suggests that the boundaries in the Abd-B regulatory domains pair with their neighbors head-to-head, and thus form circle-loops. While the contact pattern between neighbors in the Abd-B region fit with that expected for an array of circle-loop TADs, this has not been confirmed by examining the MicroC profiles before and after manipulating the boundary elements in this region. For this reason, we sought unambiguous examples of chromatin loops generated by the orientation-dependent pairing of endogenous TAD boundaries that have either a stem-loop or a circle-loop topology. The collection of meta-loops described recently by Mohana et al., 2023 provide one such test, as many appear to be generated by the pairing of TAD boundaries, and their local interaction profiles are easily interpreted.

Shown in Figure 8A is a 2.8 Mb meta-loop on chromosome 2L generated by the pairing of two TAD boundaries, labeled blue and purple (block arrows). The pairing of the blue and purple TAD boundaries brings sequences in the TADs flanking the two boundaries into contact, and this generates two rectangular boxes of interaction indicated by the arrows (see also blue double arrows in the diagram on the right). In the rectangular box on the upper left of the contact map, sequences in the TAD containing CG33543, Obp22a, and Npc2a located just upstream of the blue boundary are crosslinked to sequences upstream of the purple boundary in the TAD containing the fipi gene. In the rectangular box on the lower right, sequences in a small TAD downstream of the blue boundary (which contains Nplp4 and CG15353) are crosslinked to sequences in a TAD downstream of the purple boundary (which contains CG3294 and slf). As shown in the diagram, this pattern of interaction (upstream-to-upstream and downstream-to-downstream) indicates that the blue and purple TAD boundaries pair with each other head-to-head, and this orientation generates a large loop with a circle-loop topology.

Figure 8. Circle-loop and stem-loop meta-loops.

(A) CG3294 circle-loop meta-loop. In this meta-loop, a TAD boundary (blue arrow) located at ~2.0 Mb on chromosome 2L pairs head-to-head with a TAD boundary (purple arrow) located ~2.8 Mb away. As indicated in the diagram, head-to-head pairing generates a circle-loop. In the circle-loop topology, the TAD upstream of the blue boundary is brought into contact with the TAD upstream of the purple boundary, as indicated the diagram (blue double arrows). This generates a rectangular box of enhanced contacts between sequences in the TAD containing the CG33543, Obp22a, and Npc2a genes and sequences in a TAD that contains the fipi gene. This box is located on the upper left of the contact map (above and to the left of the black arrow). Sequences in TADs downstream of the blue and purple boundaries are also linked, and this generates a small rectangular box representing sequences in the small Nplp4 and CG15353 TAD ligated to sequences in the TAD containing CG3294 and slf (below and to the right of the black arrow). (B) The bin/Mp meta-loops on the left arm of the third chromosome are generated by the head-to-tail pairing of two sets of boundaries, indicated by the blue, brown, green, and purple arrows. Pairing of the brown and green boundaries generates an ~2.2 Mb stem-loop. Sequences in the TAD downstream of the brown boundary (which contains the Dhc64C and CG1808 genes) are linked to sequences in the TAD upstream of the green boundary (which contains CG2328 and bin). This generates the rectangular box of enhanced contacts on the lower left of the contact map. Pairing of the blue and purple boundaries head-to-tail generates a small stem-loop ‘bubble’ (see diagram). This bubble brings sequences in the TAD containing the most distal Mp promoter (blue arrowhead) into contact with sequences in the small TAD containing CG7509 (see diagram on the right). Interactions between these two TADs generates the small rectangular box of enhanced contacts in the center of the contact map. The head-to-tail pairing of the blue and purple boundaries also bring sequences in the TAD upstream of the blue boundary that contains the RhoGEF64C gene into contact with the TAD containing one of the internal Mp promoters (black arrowhead). This interaction generates the box of enhanced contacts in the upper-right portion of the contact map. The bin size for each panel is 200 bp; embryos are 12–16 hr old.

Figure 8.

Figure 8—figure supplement 1. MicroC patterns of DNA segments on the left and right arm of chromosome 2.

Figure 8—figure supplement 1.

(A) MicroC contact profiles of the left arm of chromosome 2 around 3,000,000 bp. The black arrow indicates where interactions between neighboring TADs are suppressed, as might be expected for a stem-loop configuration. The red arrow just above and to the right of the black arrow points to contacts between next-next-(next)-door neighbors that are enhanced. However, the plume that is formed is one-sided, as indicated by the green arrow. (B) MicroC contact profiles for TADs in a portion of the right arm of chromosome 2. The TADs indicated by the double-headed blue arrow at the bottom have a complex pattern of neighbor interactions. The asterisk and purple arrow indicate a potential volcano plume; however, unlike the eve volcano triangle and plume, this plume appears to be generated by crosslinking between sequences to either side of the collection of TADs indicated by the double-headed blue arrow. The TAD-to-TAD interaction patterns are further complicated by a band of enhanced crosslinking (red arrow).

The 2.2 Mb meta-loop on chromosome 3L in Figure 8B is more complicated in that it is generated by four TAD boundaries (indicated by blue, brown, green, and purple arrows). The blue and brown boundaries separate a small TAD containing CG7509 from two larger TADs, while the green and blue boundaries define the endpoints of a TAD containing the most distal promoter (blue arrowhead) of the Mp (Multiplexin) gene. As indicated in the accompanying diagram, the brown and green boundaries pair with each other head-to-tail as do the blue and purple boundaries. Pairing of the brown and green boundaries generates a large ~2.2 Mb stem-loop that brings sequences in the TAD downstream of the brown boundary, which contains the Dhc64C and CG1808 genes, into contact with sequences in the TAD upstream of the green boundary, which contains the CG3238 and bin (binou) genes. This linkage generates the rectangular box of enhanced contacts in the lower-left corner of the contact map. Pairing of the blue and purple boundaries generates a small stem-loop bubble that links sequences in the CG7509 TAD to the TAD containing the Mp distal promoter (blue arrowhead). This connection generates a small rectangular box of enhanced crosslinking in the center of the contact map. In addition, sequences upstream of the blue boundary and downstream of the purple boundary are brought into contact by the head-to-tail pairing of these two boundaries. This generates a third rectangular box of enhanced physical contact in the upper-right corner of the contact map that links sequences in the TAD containing the RhoGEF64C gene to sequences in the TAD containing the internal Mp promoter (black arrowhead). Note that the positioning of the lower-left and upper-right rectangular boxes of enhanced contact in the bin-MP meta-loop is the mirror image of the rectangular boxes of enhanced contact for the CG3294 meta-loop.

Stem-loops versus circle-loops

The results we have reported here demonstrate that boundary:boundary pairing in flies generates loops that can have either a stem-loop or a circle-loop topology. An important question is, what is the relative frequency of stem-loops versus circle-loops in the fly genome? The MicroC contact patterns for the stem-loop and circle-loop versions of the eve TAD are quite distinct. The former is a volcano triangle with a plume while the latter is a volcano triangle flanked by clouds. A survey of the MicroC contact patterns elsewhere in the non-repetitive regions of the fly genome indicates that volcanoes with plumes are rare (~30). For example, there are two volcano triangles with plumes in the Antennapedia complex, and they encompass the deformed and fushi-tarazu genes (Levo et al., 2022). However, since most of the ‘euchromatic’ regions of the fly genome are assembled into TADs whose MicroC profiles resemble that observed for eve circle-loops and the Abd-B region of BX-C, it is possible that much of the fly genome is assembled into circle-loops, not stem-loops.

While this suggestion is consistent with the available data, it is based on contact patterns between neighboring TADs, and important caveats remain. For one, the contact patterns between neighboring TADs can deviate in one way or another from that seen in the Abd-B region. For example, there are TADs in which interactions with one set of neighbors appear to be suppressed as expected for stem-loops, but the classic plume is absent, as interactions are not suppressed with the other neighbors (c.f., Figure 8—figure supplement 1A). In other cases, there is a series of complicated TAD-TAD interactions topped by a rectangular plume (Figure 8—figure supplement 1, purple arrow). For this reason, it will not be possible to draw firm conclusions about the frequency of stem-loops versus circle-loops genome-wide until the relative orientation of the paired boundaries themselves can be determined directly. On the other hand, it is clear from our studies that both classes of TADs must exist in flies. If, as seems likely, a significant fraction of the TADs genome-wide are circle-loops, this would effectively exclude cohesin-based loop extrusion as a general mechanism for TAD formation in flies. In addition, though stem-loops could be generated by a cohesin-dependent mechanism, it is unlikely that this mechanism is operational in flies, as we have shown here and in Bing et al., 2024 that stem-loops in flies are formed by orientation-dependent boundary:boundary pairing.

Another important question is whether our findings have any relevance to the formation and topology of TADs in mammals. In the most common version of the loop extrusion model, the mammalian genome is assembled into an alternating pattern of stem-loops and unanchored loops (Davidson and Peters, 2021; Higashi and Uhlmann, 2022; Perea-Resa et al., 2021). In this case, one might expect to observe volcano triangles topped by plumes alternating with DNA segments that have a considerably lower density of internal contacts. However, this crosslinking pattern is not observed in published MicroC data sets (Hsieh et al., 2020; Krietenstein et al., 2020). Instead of an alternating pattern of high-density TAD triangles separated by regions of low-density contacts, the TAD triangles are generally linked to both neighbors, just as in flies. Moreover, also like in Drosophila, there are few stem-loop volcano TADs topped by plumes. Instead, the crosslinking pattern between neighboring TADs appears similar to that observed for circle-loops in flies. Of course, one problem with these MicroC studies is that the resolution may not be sufficient to detect volcanoes with plumes or the other features predicted by the loop-extrusion model. However, there are no obvious volcanoes with plumes in the much higher resolution RCMC studies of Goel et al., 2023. Instead, the MicroC profiles most closely resemble those seen in the Abd-B region of BX-C (c.f. the Ppm1g locus in Figure 4 of Goel et al., 2023). Moreover, compromising cohesin activity has minimal impact on the TADs in this region of the mouse genome, as evidenced from the MicroC pattern before and after knockdown. Based on these observations, one can reasonably question whether cohesin-mediated loop extrusion is deployed in mammals as the mechanism for not only generating TADs but also determining TAD boundaries. Clearly, validation of the loop-extrusion/CTCF road-block model as currently formulated will require a direct demonstration that mammalian TADs are exclusively either stem-loops or unanchored loops, and that the endpoints are always (or almost always) determined by CTCF roadblocks.

TADs and A/B compartmentalization

A/B compartmentalization has been proposed as a mechanism for subdividing the chromosome into discrete domains that is independent of cohesin-mediated loop extrusion and CTCF. In this model, shared biochemical/biophysical properties that reflect the relative transcriptional state of each chromosomal segment drive block polymer co-segregation into a series of discrete domains (Rowley and Corces, 2018; Rowley et al., 2017). While previous studies suggested that the A and B compartments represented Mb-scale DNA segments, in more recent studies, Harris et al., 2023 found that the average compartment size is on the order of 12 kb. Not only is this much smaller than originally suggested, it is also similar in size to that of most TADs in the Drosophila genome, including the eve TAD. Moreover, in their studies (and also in our data sets), there is close to a one-to-one correspondence between the linear arrangement of individual TADs along the chromosome and the DNA segments that are thought to assemble into discrete domains by co-polymer segregation. This close connection to TADs is also reflected in the patchwork patterns of interacting chromatin domains that are visualized in studies on A/B compartments.

According to this newer version of the compartment model, the chromatin state of each DNA segment determines not only whether it will assemble into a discrete domain, but also how the resulting domain interacts with next-door neighbors, next-next-door neighbors, etc. However, this model does not appear to fit with several of our findings. To begin with, the sequences included in the eve TAD and its patterns of interaction with neighboring TADs are essentially the same in NC14 embryos as they are in 12–16 hr embryos. In the former case, eve is transcriptionally poised (Chen et al., 2013), probably in most or all nuclei, while in the latter case, the entire eve TAD is silenced by a PcG-dependent mechanism in all but a few nuclei (Nègre et al., 2010). However, this transition from potentially active to silenced does not impact the eve TAD, nor does it alter how the eve TAD interacts with the neighboring TADs that are (mostly) transcriptionally active at both stages of development. Similarly, in the meta-loops we have examined, the transcriptional state of the TADs and their contact patterns with their neighbors are not consistent with a strict partitioning of chromosomal segments into one of two compartments. For example, in the CG3294 meta-loop (Figure 8A), the three genes (CG33534, the odorant binding gene Obp22a and the Npc2a gene) that comprise the TAD upstream of the blue boundary would be predicted to be in the same chromatin state; however, while CG33534 and Obp22a are not expressed in embryos, the Npc2a gene is expressed at high levels during embryogenesis and should partition into a separate TAD. The TAD downstream of the blue boundary contains two transcriptionally repressed genes (CG15353 and Nplp4) in 12–16 hr embryos. This TAD physically interacts with the TAD downstream of the purple boundary that contains CG3294 and slf. According to the A/B compartment model, the CG15353 and Nplp4 TAD interacts with the CG3294 and slf TAD because the chromatin in these two TADs share biochemical/biophysical properties that are characteristic of the inactive B compartment. However, unlike CG15353 and Nplp4, both slf and CG3294 are expressed in 12–16 hr embryos, slf at a high level and CG3294 at a low level.

Likewise, if block-polymer co-segregation is the determining factor for both TAD formation and the patterns of TAD:TAD interactions, then our manipulations of the nhomie boundary should have only a minimal impact on the MicroC contact maps, unless there are significant changes in the transcriptional status of eve and its neighbors. In the lambda DNA replacement, the left endpoint of the WT eve TAD (the normal location of nhomie) in 12–16 hr embryos is lost, and instead it appears to map primarily to the right or left boundaries of the TL-3 sub-TAD. TL-3 contains the eIF3j gene, which is expressed at high levels throughout the embryo at this stage, while eve itself is silenced by a PcG-dependent mechanism in all but a small number of cells. The nhomie reverse and homie forward replacements restore the eve TAD. This means that in this instance, TAD formation is mediated by the pairing of the two replacement boundaries with homie (as demonstrated by the viewpoints in Figure 6) and not by partitioning into an A or B compartment. The replacements also alter interactions between eve and the neighboring TADs. Unlike in WT where the eve TAD is physically isolated from its neighbors, the eve TAD interacts with both neighbors in these two replacements. However, the genes in the neighboring TADs do not share the same biochemical/biophysical state—they are active in 12–16 hr embryos, and so should segregate into the A compartment, while eve is inactive and should segregate into the B compartment.

While these observations are inconsistent with a model in which block-polymer co-segregation is responsible for the formation of TADs and determining the pattern of TAD:TAD interactions, this does not rule out a different role, namely in augmenting the insulating activity of boundary elements. One of the defining properties of boundary elements is to restrict the activities of enhancers and silencers, and this helps ensure that the chromatin within a given TAD shares the same biochemical and biophysical properties. The shared biochemical/biophysical properties could in turn enhance the segregation of the chromatin into different compartments, and thus mediate some of the changes we observe. As we cannot rule out the possibility that such biophysical forces augment the functional properties of boundaries, we should add them to the list of downstream events that are dependent upon boundary–boundary interactions and the specific topologies that they can induce. So, while compartment co-segregation may well play a role, on multiple length scales, in mediating the effects of boundaries and other regulatory elements on gene expression and chromosome topology, it certainly cannot ‘replace’ their functional properties as an explanation for those effects.

Materials and methods

Creation of nhomie deletion flies

To modify nhomie at the eve locus, we used recombinase-mediated cassette exchange (Bateman et al., 2006). First, we inserted two closely positioned attP sites using CRISPR. The donor plasmid for this was constructed as follows. First, a mini-white (mw) gene with Glass binding sites (Fujioka et al., 1999) was inserted into pBlueScript. From the standard mw gene, the Wari insulator (Chetverina et al., 2008) was deleted. Then, two 102 bp attP sequences (Venken et al., 2011) were inserted, one just 5′ of the Glass binding sites and the other at the 3′ end of the modified mw, creating the plasmid pP-attPx2-mw. 5′- and 3′-homologous arms were added to both ends. Two gRNA sequences were cloned into plasmid pCFD4 (Port et al., 2014) (Addgene). The donor and gRNA plasmids were injected into a Cas9 line (y[1] M{vas-Cas9.S}ZH-2A w[1118], Bloomington Drosophila Stock Center). This chromosomal modification resulted in one attP site being inserted in the intron of CG12134, and the other being inserted between the eve 7-stripe enhancer and the 3+7 stripe enhancer. This also deleted 2.2 kb of endogenous sequence, including nhomie and the eve 7-stripe enhancer.

After identifying a successful insertion (NattPmw), mw was replaced by each of the following using RMCE: (1) the previously deleted 2.2 kb, restoring nhomie and the eve 7-stripe enhancer, to create ‘wild-type nhomie’’ (nhomie forward), (2) the same 2.2 kb sequence, but with 600 bp of phage λ DNA in place of 600 bp nhomie (λ DNA), (3) the same 2.2 kb sequence, but with 600 bp nhomie inverted (nhomie reverse), and (4) the same 2.2 kb sequence, but with 600 bp nhomie replaced by a copy of ~600 bp homie in its native orientation in the chromosome (homie forward). Each of these changes was confirmed by sequencing of genomic DNA from the transgenic fly lines.

Analysis of embryonic cuticle patterns and in situ hybridization

To identify defects in developing embryos, embryos were collected for 2.5 hr, and allowed to develop for an additional 20–21 hr at 25°C. Embryos were dechorionated and mounted in a 1:1 mixture of Hoyer’s medium and lactic acid. Mounted embryos were left at room temperature (RT) until they cleared (12–14 days), and the patterns of ventral abdominal denticles were examined and tallied as follows. Loss of at least one-fifth of a denticle band (in A1-A8) was counted as ‘missing’. Fused denticle bands, which rarely occurred, were also counted as a ‘missing’ band. Minor defects such as those within individual denticle rows were not counted.

Digoxigenin (DIG) in situ hybridization was performed using DIG-labeled anti-sense RNA against CG12134, eIF3j, and TER94. RNA expression was visualized using alkaline phosphatase-conjugated anti-DIG antibody (Roche), using CBIP and NBT as substrates (Roche). Each set of experiments was carried out in parallel to minimize experimental variation. Representative expression patterns are shown in each figure.

HCR-FISH

The sequences of target genes were obtained from FlyBase (https://www.flybase.org/; Gramates et al., 2022). To design probes, the target gene sequences were submitted to the Molecular Instruments probe design platform (https://molecularinstruments.com/hcr-rnafish; Choi et al., 2016), with parameters set to a 35 probe set size for Drosophila melanogaster. A similar method was designed based on published smFISH methods (Little and Gregor, 2018; Trcek et al., 2017). 100–200 flies were placed in a cage with an apple juice plate at the bottom. For early stages, the embryos were collected for 7 hr, while for later-stage embryos, collections were overnight. Embryos from each plate were washed into collection mesh and dechorionated in bleach for 2 min, then fixed in 5 mL of 4% paraformaldehyde in 1× PBS and 5 mL of heptane for 15 min with horizontal shaking. The paraformaldehyde was then removed and replaced with 5 mL methanol. The embryos were then devitellinized by vortexing for 30 s and washed in 1 mL of methanol twice. Methanol was then removed and replaced by PTw (1× PBS with 0.1% Tween-20) through serial dilution as 7:3, 1:1, and 3:7 methanol:PTw. The embryos were washed twice in 1 mL of PTw and pre-hybridized in 200 μL of probe hybridization buffer for 30 min at 37°C. 0.4 pmol of each probe set were added to the embryos in the probe hybridization buffer, and the embryos were incubated at 37°C for 12–14 hr. The embryos were then washed 3× with probe wash buffer at 37°C for 30 min and 2× with 5× SSCT (5× SSC + 0.1% tween) at RT for 5 min. Then the embryos were pre-amplified with 300 μL amplification buffer for 10 min at 25°C. Meanwhile, 6 pmol of hairpin h1 and h2 were snap-cooled separately (95°C for 90 s, cool to RT with 0.1°C drop per second), and then mixed in 100 μL of amplification buffer at RT. After that, the pre-amplification solution was removed from the embryos, and 100 μL of hairpin h1/h2 mix were added to the embryos. Next, the embryos were incubated for 12–14 hr at RT in the dark. To remove excess hairpins, the embryos were washed in SSCT as follows: 2× for 5 min, 2× for 30 min, and 5× for 5 min. Then, the embryos were washed with 1 mL PTw for 2 min and stained with DAPI/Hoechst at 1 μg/mL for 15 min at RT in the dark. The embryos were then washed with PTw 3× for 5 min. Finally, the embryos were mounted on microscope slides with Vectashield and a #1.5 coverslip for imaging.

Imaging, image analysis, and statistics

Embryos from HCR-FISH were imaged using a Nikon A1 confocal microscope system, with a Plan Apo ×20/0.75 DIC objective. Z-stack images were taken at interval of 2 μm, 4× average, 1024 × 1024 resolution, and the appropriate laser power and gain were set for 405, 488, 561, and 640 channels to avoid overexposure. Images were processed using ImageJ, and the maximum projection was applied to each of the stack images. To determine the presence of stripes in early embryos, multi-channel images were first split into single channels and the stripe signal was highlighted and detected by the MaxEntropy thresholding method. GraphPad Prism was used for data visualization and statistical analysis. Two-way ANOVA with Tukey’s multiple comparisons test for each pair of groups was used to determine the statistical significance for the percentage of embryos carrying stripes in eIF3j and TER94 channels in each group.

MicroC library construction for the nhomie replacements

Embryos were collected on yeasted apple juice plates in population cages for 4 hr, incubated for 12 hr at 25°C, then subjected to fixation as follows. Embryos were dechorionated for 2 min in 3% sodium hypochlorite, rinsed with deionized water, and transferred to glass vials containing 5 mL PBST (0.1% Triton-X100 in PBS), 7.5 mL n-heptane, and 1.5 mL fresh 16% formaldehyde. Crosslinking was carried out at RT for exactly 15 min on an orbital shaker at 250 rpm, followed by addition of 3.7 mL 2 M Tris-HCl pH 7.5 and shaking for 5 min to quench the reaction. Embryos were washed twice with 15 mL PBST and subjected to secondary crosslinking. Secondary crosslinking was done in 10 mL of freshly prepared 3 mM final DSG and ESG in PBST for 45 min at RT with passive mixing. The reaction was quenched by addition of 3.7 mL of 2 M Tris-HCl pH7.5 for 5 min, washed twice with PBST, snap-frozen, and stored at –80°C until library construction.

Micro-C libraries were prepared as previously described (Batut et al., 2022) with the following modifications: 50 µL of 12–16 hr embryos were used for each biological replicate. 60U of MNase was used for each reaction to digest chromatin to a mononucleosome:dinucleosome ratio of 4. Libraries were barcoded, pooled, and subjected to paired-end sequencing on an Illumina Novaseq S1 100nt Flowcell (read length 50 bases per mate, 6-base index read).

Two or more independent biological replicates were sequenced for each genotype. For each replicate, >1000 embryos were used, and ~250M reads sequenced. Post-sequencing QC analysis was done for every sample, and the QC reports are available along with the sequence data in GEO (GSE263270). The raw sequencing data are also available in GSE263270 for use in further bioinformatics analysis. The figures present the merged data from all independent biological replicates for each genotype. The total read numbers in the merged data are very similar for each genotype (~500M reads). For NC14 embryo MicroC (Figure 2), public data sets GSE171396 and GSE173518 were used (Batut et al., 2022; Levo et al., 2022).

MicroC data processing

MicroC data for D. melanogaster were aligned to custom genomes edited from the Berkeley Drosophila Genome Project (BDGP) Release 6 reference assembly (dos Santos et al., 2015) with BWA-MEM (Li and Durbin, 2009) using parameters -S -P -5 -M. The resultant BAM files were parsed, sorted, de-duplicated, filtered, and split with Pairtools (https://github.com/open2c/pairtools; Goloborodko, 2024). We removed pairs where only half of the pair could be mapped, or where the MAPQ score was less than three. The resultant files were indexed with Pairix (https://github.com/4dn-dcic/pairix; Lee, 2024). The files from replicates were merged with Pairtools before generating 100 bp contact matrices using Cooler (Abdennur and Mirny, 2020). Finally, balancing and Mcool file generation were performed with Cooler’s Zoomify tool.

Virtual 4C profiles were extracted from individual replicates using FAN-C (Kruse et al., 2020) at 400 bp resolution. The values were summed across replicates and smoothed across three bins (1.2 kb). The homie viewpoint was set to the 549nt homie sequence that was defined in previous studies (Fujioka et al., 2016; Fujioka et al., 2009).

Acknowledgements

We thank Gordon Grey for running the fly food facility at Princeton, members of the Lewis Sigler Genomics Core facility for their invaluable assistance with DNA sequencing, and Qing Liu for excellent technical assistance. We would also like to thank members of MOL431 for creative input. Special thanks to Olga Kyrchanova, Daria Chetverina, Maksim Erokhin, Pavel Georigev, Tsutomu Aoki, Girish Deshpande, Airat Ibragimov, Sergey Ryabichko, Yuri Pritykin, Alex Ostrin, Xinyang Bing, Xiao Li, and Mike Levine for stimulating discussions and sharing unpublished data.

Appendix 1

Appendix 1—key resources table.

Reagent type (species) or resource Designation Source or reference Identifiers Additional information
Gene (Drosophila melanogaster) eve FlyBase FBgn0000606
Gene (D. melanogaster) CG12134 FlyBase FBgn0033471
Gene (D. melanogaster) eIF3j FlyBase FBgn0027619
Gene (D. melanogaster) TER94 FlyBase FBgn0286784
Genetic reagent (D. melanogaster) y1 M{vas-Cas9}ZH-2A w1118/FM7c Bloomington Drosophila Stock Center 51323
Recombinant DNA reagent
(plasmid)
pCFD4-U6:1_U6:3tandemgRNAs Addgene 49411
Chemical compound, drug n-Heptane Fisher Chemical O3008-4
Chemical compound, drug Paraformaldehyde 20% solution, EM Grade Electron Microscopy Sciences 15713S
Chemical compound, drug Formaldehyde, 16%, methanol free, Ultra Pure Polysciences Inc 18814-10
Chemical compound, drug PBS – phosphate-buffered saline (10×) pH 7.4, RNase-free Thermo Fisher AM9624
Chemical compound, drug Tween 20 Sigma P1379
Chemical compound, drug Triton X-100 Bio-Rad 161-0407
Chemical compound, drug Tris base Sigma 11814273001
Chemical compound, drug Methanol Fisher Chemical 203403
Chemical compound, drug SSC, 20× Thermo Fisher 15557044
Chemical compound, drug Formamide Thermo Fisher 17899
Chemical compound, drug Dextran sulfate Sigma D8906
Chemical compound, drug Salmon Sperm DNA Thermo Fisher AM9680
Chemical compound, drug Ribonucleoside Vanadyl Complex NEB S1402S
Chemical compound, drug Nuclease-free BSA Sigma 126609
Chemical compound, drug Triethylammonium acetate Sigma 625718
Chemical compound, drug dGTP (100 MM) VWR 76510-208
Chemical compound, drug dTTP (100 MM) VWR 76510-224
Chemical compound, drug Lonza NuSieve 3:1 Agarose Thermo Fisher BMA50090
Other T4 DNA ligase NEB M0202L Enzyme
Chemical compound, drug Biotin-11-dCTP Jena Bioscience NU-809-BIOX
Chemical compound, drug Biotin-14-dATP Jena Bioscience NU-835-BIO14
Commercial assay or kit Qubit dsDNA HS Assay Kit Life Technologies Corp. Q32851
Chemical compound, drug Atto 633 NHS ester Sigma 01464
Chemical compound, drug Phase Lock Gel, QuantaBio - 2302830, Phase Lock Gel Heavy VMR 10847-802
Commercial assay or kit NEBNext Ultra II DNA Library Prep Kit for Illumina NEB E7645S
Commercial assay or kit Ampure Xp 5 ml Kit Thermo Fisher NC9959336
Commercial assay or kit Hifi Hotstart Ready Mix Thermo Fisher 501965217
Commercial assay or kit Dynabeads MyOne Streptavidin C1 Life Technologies Corp. 65001
Chemical compound, drug cOmplete, EDTA-free Protease Inhibitor Cocktail Sigma 11873580001
Chemical compound, drug N,N-Dimethylformamide Sigma 227056
Chemical compound, drug Potassium acetate solution Sigma 95843
Chemical compound, drug DSG (disuccinimidyl glutarate) Thermo Fisher PI20593
Other T4 Polynucleotide Kinase – 500 units NEB M0201S Enzyme
Other DNA Polymerase I, Large (Klenow) Fragment – 1000 units NEB M0210L Enzyme
Commercial assay or kit End-it DNA End Repair Kit Thermo Fisher NC0105678
Other Proteinase K recomb. 100 mg Sigma 3115879001 Enzyme
Other Nuclease Micrococcal (s7) Thermo Fisher NC9391488 Enzyme
Chemical compound, drug EGS (ethylene glycol bis(succinimidyl succinate)) Thermo Fisher PI21565
Commercial assay or kit Atto 565 NHS ester Sigma 72464
Commercial assay or kit HCR RNA-FISH Custom Probe Set: eve Molecular Instruments Custom probes
Commercial assay or kit HCR RNA-FISH Custom Probe Set: ter94 Molecular Instruments Custom probes
Commercial assay or kit HCR RNA-FISH Custom Probe Set: CG12134 Molecular Instruments Custom probes
Commercial assay or kit HCR RNA-FISH Custom Probe Set: eIF3j Molecular Instruments Custom probes
Commercial assay or kit HCR Amplifier B1, 488 Molecular Instruments Custom probes
Commercial assay or kit HCR Amplifier B2, 564 Molecular Instruments Custom probes
Commercial assay or kit HCR Amplifier B3, 647 Molecular Instruments Custom probes
Commercial assay or kit HCR Buffers Molecular Instruments Custom probes
Commercial assay or kit NEBNext Multiplex Oligos for Illumina NEB E7335S
Software, algorithm Fiji (ImageJ) Schindelin et al., 2012 fiji.sc
Software, algorithm NIS element Nikon microscope.healthcare.nikon.com/products/software/nis-elements
Software, algorithm GraphPad Prism 8 GraphPad Software https://www.graphpad.com/
Software, algorithm HiGlass Kerpedjiev et al., 2018 https://higlass.io/app
Software, algorithm bwa Li and Durbin, 2009 https://bio-bwa.sourceforge.net/
Software, algorithm samtools GitHub/open source https://samtools.github.io
Software, algorithm pairsamtools Goloborodko et al., 2024 https://github.com/mirnylab/pairsamtools
Software, algorithm pairix Lee, 2024 https://github.com/4dn-dcic/pairix
Software, algorithm cooler Abdennur and Mirny, 2020; Abdennur, 2016 https://github.com/open2c/cooler
Software, algorithm Miniconda Anaconda https;//docs.conda.io/en/latest/miniconda/
Software, algorithm Snakemake GitHub/open source https://snakemake.github.io

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Contributor Information

Paul Schedl, Email: pschedl@princeton.edu.

James B Jaynes, Email: james.jaynes@jefferson.edu.

Yukiko M Yamashita, Whitehead Institute/MIT, United States.

Claude Desplan, New York University, United States.

Funding Information

This paper was supported by the following grants:

  • New Jersey Commission on Cancer Research COCR23PDF011 to Wenfan Ke.

  • Histochemical Society Keystone Grant to Wenfan Ke.

  • National Institute of General Medical Sciences R35 GM126975 to Paul Schedl.

  • National Institute of General Medical Sciences R01 GM137062 to James B Jaynes.

Additional information

Competing interests

No competing interests declared.

Author contributions

Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology.

Conceptualization, Formal analysis, Investigation, Methodology.

Conceptualization, Supervision, Funding acquisition, Writing - original draft, Writing - review and editing.

Conceptualization, Supervision, Funding acquisition, Writing - review and editing.

Additional files

MDAR checklist

Data availability

Sequence data are available at GEO GSE263270. Confocal images are available on Open Science Framework at https://doi.org/10.17605/OSF.IO/6PYBM.

The following datasets were generated:

Ke W, Fujioka M, Schedl P, Jaynes J. 2024. Chromosome Structure II: Stem-loops and circle-loops. NCBI Gene Expression Omnibus. GSE263270

Ke W. 2024. Stem-loop and circle-loop TADs generated by directional pairing of boundary elements have distinct physical and regulatory properties. Open Science Framework. 6pybm/

The following previously published datasets were used:

Bing X, Batut P, Levine M. 2022. Genome organization controls transcriptional dynamics during development. NCBI Gene Expression Omnibus. GSE171396

Bing X, Levo M, Raimundo J, Levine M. 2022. Transcriptional coupling of distant regulatory genes in living embryos. NCBI Gene Expression Omnibus. GSE173518

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eLife assessment

Yukiko M Yamashita 1

This valuable work investigates the role of boundary elements in the formation of 3D genome architecture. The authors established a specific model system that allowed them to manipulate boundary elements and examine the resulting genome topology. The work yielded the first demonstration of the existence of stem and circle loops in a genome and confirms a model which had been posited based on extensive prior genetic work, providing insights into how 3D genome topologies affect enhancer–promoter communication. The evidence is solid, although the degree of generalization remains uncertain.

Reviewer #1 (Public Review):

Anonymous

In this study, the authors engineer the endogenous left boundary of the Drosophila eve TAD, replacing the endogenous Nhomie boundary by either a neutral DNA, a wildtype Nhomie boundary, an inverted Nhomie boundary, or a second copy of the Homie boundary. They perform Micro-C on young embryos and conclude that endogenous Nhomie and Homie boundaries flanking eve pair with head-to-tail directionality to form a chromosomal stem loop. Abrogating the Nhomie boundary leads to ectopic activation of genes in the former neighboring TAD by eve embryonic stripe enhancers. Replacing Nhomie by an inverted version or by Homie (which pairs with itself head-to-head) transformed the stem loop into a circle loop. An important finding was that stem and circle loops differentially impact endogenous gene regulation both within the eve TAD and in the TADs bracketing eve. Intriguingly, an eve TAD with a circle loop configuration leads to ectopic activation of flanking genes by eve enhancers - indicating compromised regulatory boundary activity despite the presence of an eve TAD with intact left and right boundaries.

The results obtained are of high-quality and are meticulously discussed. This work advances our fundamental understanding of how 3D genome topologies affect enhancer-promoter communication.

This study raises interesting questions to be addressed in future studies.

First, given the unique specificity with which Nhomie and Homie pair (and exhibit "homing" activity), the generalizability of TAD formation by directional boundary pairing remains unclear. Testing whether boundary pairing is a phenomenon restricted to exceptional loci picked for study, rather than a broader rule of TAD formation, would best be done through the development of untargeted approaches to study boundary pairing.

Second, boundary pairing is one of several mechanisms that may form chromosomal contact domains such as TADs. Other mechanisms include cohesin-mediated chromosomal loop extrusion and the inherent tendency of transcriptionally active and inactive chromatin to segregate (or compartmentalize). The functional interplay between these possible TAD-forming mechanisms remains to be further investigated.

Reviewer #2 (Public Review):

Anonymous

This study reports a set of experiments and subsequent analyses focusing on the role of Drosophila boundary elements in shaping 3D genome structure and regulating gene expression. The authors primarily focus on the region of the fly genome containing the even skipped (eve) gene; eve is expressed in a canonical spatial pattern in fly embryos and its locus is flanked by the well-characterized neighbor of homie (nhomie) and homie boundary elements. The main focus of the investigation is the orientation dependence of these boundary elements, which had been observed previously using reporter assays. In this study, the authors use Crispr/Cas9 editing followed by recombination-mediated cassette exchange to create a series of recombinant fly lines in which the nhomie boundary element is either replaced with exongenous sequence from phage 𝝀, an inversion of nhomie, or a copy of homie that has the same orientation as the endogenous homie sequence. The nhomie sequence is also regenerated in its native orientation to control for effects introduced by the transgenesis process.

The authors then perform high-resolution Micro-C to analyze 3D structure and couple this with fluorescent and colorimetric RNA in situ hybridization experiments to measure the expression of eve and nearby genes during different stages of fly development. The major findings of these experiments are that total loss of boundary sequence (replacement with 𝝀 DNA) results in major 3D structure changes and the most prominent observed gene changes, while inversion of the nhomie boundary or replacement with homie resulted in more modest effects in terms of 3D structure and gene expression changes and a distinct pattern of gene expression change from the 𝝀 DNA replacement. As the samples in which the nhomie boundary is inverted or replaced with homie have similar Micro-C profiles at the eve locus and show similar patterns of a spurious gene activation relative to the control, the observed effects appear to be driven by the relative orientation of the nhomie and homie boundary elements to one another.

Collectively, the findings reported in the manuscript are of broad interest to the 3D genome field. Although extensive work has gone into characterizing the patterns of 3D genome organization in a whole host of species, the underlying mechanisms that structure genomes and their functional consequences are still poorly understood. The perhaps best understood system, mechanistically, is the coordinated action of CTCF with the cohesin complex, which in vertebrates appears to shape 3D contact maps through a loop extrusion-pausing mechanism that relies on orientation-dependent sequence elements found at the boundaries of interacting chromatin loops. Despite having a CTCF paralog and cohesin, the Drosophila genome does not appear to be structured by loop extrusion-pausing. The identification of orientation-dependent elements with pronounced structural effects on genome folding thus may shed light on alternative mechanisms used to regulated genome structure, which in turn may yield insights into the significance of particular folding patterns.

On the whole, this study is comprehensive and represents a useful contribution to the 3D genome field. The transgenic lines and Micro-C datasets generated in the course of the work will be valuable resources for the research community. Moreover, the manuscript, while dense in places, is generally clearly written and comprehensive in its description of the work. However, I have a number of comments and critiques of the manuscript, mainly centering on the framing of the experiments and presentation of the Micro-C results and on the manner in which the data are analyzed and reported.

As this document now reflects my review of a revised version of the initial preprint, I will begin to add the new content at this point. As discussed in detail in the following paragraphs, my initial impression of the manuscript has not changed, so I have accordingly left the above text unaltered.

In my initial review, I provided a number of suggestions to improve the quality of the manuscript. These suggestions, which took the form of six major and three minor points, largely focused on (1) altering the writing in certain places to make the story more broadly accessible to the readership and (2) the inclusion of key, missing methodological detail to increase the rigor and reproducibility of the study. No new experiments were requested, and all of the points could be readily addressed with rather straightforward textual changes.

In their revised manuscript, the authors elected to directly address one of the major points and two of the minor points (major point 4, minor points 1 and 3). The remainder of my suggestions remain entirely unaddressed. A similar level of responsiveness was afforded to the very reasonable critiques of the other Reviewer and the Reviewing Editor. The authors have instead largely chosen to respond to the points raised exclusively in the rebuttal document. This document sprawls across >22 pages, includes numerous in-line figures, and cites dozens of references. The tone of this document, in many places, is at best forceful. In a less generous interpretation, many sections are combative, dismissive, and borderline unprofessional.

It is my opinion that the authors are doing the scientific community a disservice with their response. While it is my understanding that readers will be able see the rebuttal letter, I find that end result far from satisfying. How many readers will take the trouble to access that file, versus the manuscript itself? Skirting the review critiques places an unfair burden on readers, who are expecting peer-reviewed science, to dig into the accessory files to follow the critique and response, rather than seeing in reflected in the final product as they accustomed. Intentionally or not, the tactics the authors have chosen detract from what is otherwise a novel and well-intentioned new publishing model. It is also worth pointing out that peer review is done as an act of service to the scientific community, as the senior authors are doubtless aware. The other reviewer, the Reviewing Editor, and I have all taken time away from advancing our own careers and those of our trainees to offer the thoughtful critiques that were so pointedly dismissed.

In summary, as the vast majority of my critiques remain unaddressed, I have simply reproduced them below.

Major Points:

(1) The authors motivate much of the introduction and results with hypothetical "stem loop" and "circle loop" models of chromosome confirmation, which they argue are reflected in the Micro-C data and help to explain the observed ISH patterns. While such structures may possibly form, the support for these specific models vs. the many alternatives is not in any way justified. For instance, no consideration is given to important biophysical properties such as persistence length, packing/scaling, and conformational entropy. As the biophysical properties of chromatin are a very trafficked topic both in terms of experimentation and computational modeling and generally considered in the analysis of chromosome conformation data, the study would be strengthened by acknowledgement of this body of work and more direct integration of its findings.

(2) Similar to Point 1, while there is a fair amount of discussion of how the observed results are or are not consistent with loop extrusion, there is no discussion of the biophysical forces that are thought to underly compartmentalization such as block-polymer co-segregation and their potential influence. I found this absence surprising, as it is generally accepted that A/B compartmentalization essentially can explain the contact maps observed in Drosophila and other non-vertebrate eukaryotes (Rowley, ..., Corces 2017; PMID 28826674). The manuscript would be strengthened by consideration of this phenomenon.

(3) The contact maps presented in the study represent many cells and distinct cell types. It is clear from single-cell Hi-C and multiplexed FISH experiments that chromosome conformation is highly variable even within populations of the same cell, let alone between cell types, with structures such as TADs being entirely absent at the single cell level and only appearing upon pseudobulking. It is difficult to square these observations with the models of relatively static structures depicted here. The authors should provide commentary on this point.

(4) Related to Point 4, the lack of quantitative details about the Micro-C data make it difficult to evaluate if the changes observed are due to biological or technical factors. It is essential that the authors provide quantitative means of controlling for factors like sampling depth, normalization, and data quality between the samples.

(5) The ISH effects reported are modest, especially in the case of the HCR. The details provided for how the imaging data were acquired and analyzed are minimal, which makes evaluating them challenging. It would strengthen the study to provide much more detail about the acquisition and analysis and to include depiction of intermediates in the analysis process, e.g. the showing segmentation of stripes.

eLife. 2024 Aug 7;13:RP94114. doi: 10.7554/eLife.94114.3.sa3

Author response

Wenfan Ke 1, Miki Fujioka 2, Paul Schedl 3, James B Jaynes 4

The following is the authors’ response to the original reviews.

Reviewer #1 (Public Review):

Summary:

In this study, the authors engineer the endogenous left boundary of the Drosophila eve TAD, replacing the endogenous Nhomie boundary by either a neutral DNA, a wildtype Nhomie boundary, an inverted Nhomie boundary, or a second copy of the Homie boundary. They perform Micro-C on young embryos and conclude that endogenous Nhomie and Homie boundaries flanking eve pair with head-to-tail directionality to form a chromosomal stem loop. Abrogating the Nhomie boundary leads to ectopic activation of genes in the former neighboring TAD by eve embryonic stripe enhancers. Replacing Nhomie by an inverted version or by Homie (which pairs with itself head-to-head) transformed the stem loop into a circle loop. An important finding was that stem and circle loops differentially impact endogenous gene regulation both within the eve TAD and in the TADs bracketing eve. Intriguingly, an eve TAD with a circle loop configuration leads to ectopic activation of flanking genes by eve enhancers - indicating compromised regulatory boundary activity despite the presence of an eve TAD with intact left and right boundaries.

Strengths:

Overall, the results obtained are of high-quality and are meticulously discussed. This work advances our fundamental understanding of how 3D genome topologies affect enhancer-promoter communication.

Weaknesses:

Though convincingly demonstrated at eve, the generalizability of TAD formation by directional boundary pairing remains unclear, though the authors propose this mechanism could underly the formation of all TADs in Drosophila and possibly even in mammals. Strong and ample evidence has been obtained to date that cohesin-mediated chromosomal loop extrusion explains the formation of a large fraction of TADs in mammals.

(1.1) The difficulty with most all of the studies on mammal TADs, cohesin, and CTCF roadblocks is that the sequencing depth is not sufficient, and large bin sizes (>1 kb) are needed to visualize chromosome architecture. The resulting contact profiles show TAD neighborhoods, not actual TADs.

This problem is illustrated by comparing the contact profiles of mammalian MicroC data sets at different bin sizes in Author response image 1. In this figure, the darkness of the “pixels” in panels E, F, G, and H was enhanced by reducing brightness using Photoshop.

Author response image 1. Mammalian MicroC profiles at different bin sizes.

Author response image 1.

[Author response image 1 is adapted from Krietenstein et al., 2020; Hsieh et al., 2020.]

Panels A and C are from Krietenstein et al. (2020), and show “TADs” using bin sizes typical of most mammalian studies. At this level of resolution, TADs, the “trees” that are the building blocks of chromosomes, are not visible. Instead, what is seen are TAD neighborhoods or “forests”. Each neighborhood consists of several dozen individual TADs. The large bins in these panels also artificially accentuated TAD:TAD interactions, generating a series of “stripes” and “dots” that correspond to TADs bumping into each other and sequences getting crosslinked. For example, in panel A there is a prominent stripe on the edge of a “TAD” (blue arrow). In panel C, this stripe resolves into a series of dots arranged as parallel, but interrupted, “stripes” (green and blue arrows). At the next level of resolution, it can be seen that the stripe marked by the blue arrow and magenta asterisk is generated by contacts between the left boundary of the TAD indicated by the magenta bar with sequences in a TAD (blue bar) ~180 kb way. While dots and stripes are prominent features in contact profiles visualized with larger bin sizes (A and C), the actual TADs that are observed with a bin size of 200 bp (examples are underlined by black bars in panel G) are not bordered by stripes, nor are they topped by obvious dots. The one possible exception is the dot that appears at the top of the volcano triangle underlined with magenta.

The chromosome 1 DNA segment from the MicroC data of Hsieh et al. (2020) shows a putative volcano triangle with a plume (indicated by a V in Author response image 1, panels D, F, and H). Sequences in the V TAD don’t crosslink with their immediate neighbors, and this gives a “plume” above the volcano triangle, as indicate by the light blue asterisk in panels D, F, and H. Interestingly, the V TAD does contact two distant TADs, U on the left and W on the right. The U TAD is ~550 kb from V, and the region of contact is indicated by the black arrow. The W TAD is ~585 kb from V, and the region of contact is indicated by the magenta arrow. While the plume still seems to be visible with a bin size of 400 bp (light blue asterisk), it is hard to discern when the bin size is 200 bp, as there are not enough reads.

The evidence demonstrating that cohesin is required for TAD formation/maintenance is based on low resolution Hi-C data, and the effects that are observed are on TAD neighborhoods (forests), and not TADs (trees). In fact, there is published evidence that cohesin is not required in mammals for TAD formation/maintenance. Author response image 2 shows the Ppm1g region of mouse chromosome 5 generated from data in a paper by Goel et al. (2023). In this experiment, the authors depleted the cohesin component RAD21 and then visualized the effects on TAD organization using the high resolution region capture MicroC (RCMC) protocol. The MicroC contact map in Author response image 2 visualizes a ~150 kb DNA segment around the Ppm1pg locus at 250 bp resolution. On the right side of the diagonal is the untreated control, while the left side shows the MicroC profile of the same region after RAD21 depletion. The authors indicated that there was a 97% depletion of RAD21 in their experiment. However, as is evident from a comparison of the experimental and control, loss of RAD21 has no apparent effect on the TAD organization of this mammalian DNA segment. Likewise, TAD:TAD interactions between next door neighbors (purple asterisks), next-next door neighbors (red asterisks) and next-next-next door neighbors (blue asterisks) is not perturbed either.

Author response image 2.

Author response image 2.

[Author response image 2 is generated from data available from Goel et al., 2023.]

Several other features are worth noting. First, unlike the MicroC experiments shown in Author response image 1, there are dots at the apex of the TADs in this chromosomal segment. In the MicroC protocol, fixed chromatin is digested to mononucleosomes by extensive MNase digestion. The resulting DNA fragments are then ligated, and dinucleosome-length fragments are isolated and sequenced. DNA sequences that are nucleosome free in chromatin (which would be promoters, enhancers, silencers and boundary elements) are typically digested to oligonucleotides in this procedure and won’t be recovered. This means that the dots shown here must correspond to mononucleosome-length elements that are MNase resistant. This is also true for the dots in the MicroC contact profiles of the Drosophila Abd-B regulatory domain (see Fig. 2B in the paper). Second, the TADs are connected to each other by 45o stripes (see blue and green arrowheads). While it is not clear from this experiment whether the stipes are generated by an active mechanism (enzyme) or by some “passive” mechanism (e.g., sliding), the stripes in this chromosomal segment are not generated by cohesin, as they are unperturbed by RAD21 depletion. Third, there are no volcano triangles with plumes in this chromosomal DNA segment. Instead, the contact patterns between neighboring TADs closely resemble those seen for the Abd-B regulatory domains (compare Author response image 2 with Fig. 2B in the paper). This similarity suggests that the TADs in and around Ppm1g may be circle-loops, not stem-loops. As volcano triangles with plumes also seem to be rare in the MicroC data sets of Krietenstein et al. (2020) and Hsieh et al. (2020) (with the caveat that these data sets are low resolution: see Author response image 1), it is possible that much of the mammalian genome is assembled into circle-loop TADs, a topology that can’t be generated by the cohesin loop-extrusion (bolo tie clip) /CTCF roadblock model.

Author response image 3.

Author response image 3.

[Author response image 3 is generated from data available from Goel et al., 2023.]

While RAD21 depletion has no apparent effect on TADs, it does appear to have a modest impact on TAD neighborhoods. This was shown in a supplemental figure in Goel et al. (2023), which visualized the Ppm1g region of chromosome 5 with bin sizes of 5 kb and 1 kb. Author response image 3 shows an ~600 kb region from chromosome 5 containing the Ppm1g gene, visualized with a bin size of 1 kb. As can be seen from comparing the MicroC profiles in this image with that in Author response image 2, individual TADs are not visible. Instead, the individual TADs are binned into large TAD “neighborhoods” that consist of multiple TADs.

Unlike the individual TADs shown in Author response image 2, the TAD neighborhoods in Author response image 3 show a limited sensitivity to RAD21 depletion. The effects of RAD21 depletion can be seen by comparing the relative pixel density in the box before (blue box above the diagonal) and after auxin-induced RAD21 degradation (purple box below the diagonal). The reduction in pixel density is greatest for more distant TAD:TAD contacts (farthest from the diagonal: green double arrow). By contrast, the TADs themselves are unaffected, as are contacts between individual TADs and their neighbors (Author response image 2 above). A subset of higher density contact “dots” (green asterisks) also appear to be reduced after RAD21 depletion, though the effects are not uniform (blue asterisks). At this point it isn’t clear why contacts between distant TADs in the same neighborhood are lost when RAD21 is depleted; however, a plausible speculation is that it is related to the functioning of cohesin in holding newly replicated DNAs together until mitosis, and whatever other role(s) it might have in chromosome condensation.

Moreover, given the unique specificity with which Nhomie and Homie are known to pair (and exhibit "homing" activity), it is conceivable that formation of the eve TAD by boundary pairing represents a phenomenon observed at exceptional loci rather than a universal rule of TAD formation. Indeed, characteristic Micro-C features of the eve TAD are only observed at a restricted number of loci in the fly genome…..

(1.2) The available evidence does not support the claim that nhomie and homie are “exceptional.” To begin with, nhomie and homie rely on precisely the same set of factors that have been implicated in the functioning of other boundaries in the fly genome. For example, homie requires (among other factors) the generic boundary protein Su(Hw) for insulation and long-distance interactions (Fujioka et al. 2024). (This is also true of nhomie: unpublished data.) The Su(Hw) protein (like other fly polydactyl zinc finger proteins) can engage in distant interactions. This was first shown by Sigrist and Pirrotta (Sigrist and Pirrotta 1997), who found that the su(Hw) element from the gypsy transposon can mediate long-distance regulatory interactions (PRE-dependent silencing) between transgenes inserted at different sites on homologous chromosomes (trans interactions) and at sites on different chromosomes.

The ability to mediate long-distance interactions is not unique to the su(Hw) element, or homie and nhomie. Muller et al. (1999) found that the Mcp boundary from the Drosophila BX-C is also able to engage in long-distance regulatory interactions: both PRE-dependent silencing of mini-white and enhancer activation of mini-white and yellow. The functioning of the Mcp boundary depends upon two other generic insulator proteins, Pita and the fly CTCF homolog (Kyrchanova et al. 2017). Like Su(Hw), both are polydactyl zinc finger proteins, and they resemble the mammalian CTCF protein in that their N-terminal domain mediates multimerization (Bonchuk et al. 2020; Zolotarev et al. 2016). Author response image 4 shows PRE-dependent “pairing sensitive silencing” interactions between transgenes carrying a mini-white reporter, the Mcp and scs’ (BEAF-dependent, Hart et al. 1997) boundary elements, and a PRE closely linked to Mcp. In this experiment, flies homozygous for different transgene inserts were mated and the eye color was examined in their trans-heterozygous progeny. As indicated in the figure, the strongest trans-silencing interactions were observed for inserts on the same chromosomal arm; however, transgenes inserted on the left arm of chromosome 3 can interact across the centromere with transgenes inserted on the right arm of chromosome 3.

Author response image 4.

Author response image 4.

[Author response image 4 is reproduced from Figure 6 from Muller et al., 1999, with permission from Genetics Society of America. It is not covered by the CC-BY 4.0 license, and further reproduction of this figure would need permission from the copyright holder.]

Author response image 5A shows a trans-silencing interaction between w#11.102 at 84D and w#11.16 approximately 5.8 Mb away, at 87D. Author response image 5B shows a trans-silencing interaction across the centromere between w#14.29 on the left arm of chromosome 3 at 78F and w#11.102 on the right arm of chromosome 3 at 84D. The eye color phenotype of mini-white-containing transgenes is usually additive: homozygyous inserts have twice as dark eye color as the corresponding hemizygous inserts. Likewise, in flies trans-heterozygous for mini-white transgenes inserted at different sites, the eye color is equivalent to the sum of the two transgenes. This is not true when mini-white transgenes are silenced by PREs. In the combination shown in panel A, the trans-heterozygous fly has a lighter eye color than either of the parents. In the combination in panel B, the trans-heterozygous fly is slightly lighter than either parent.

Author response image 5. Long-distance pairing-sensitive silencing between transgenes inserted on 3R.

Author response image 5.

Transgene insertion sites are shown in Author response image 4.

[Author response image 5 is reproduced from Figure 5C from Muller et al., 1999, with permission from Genetics Society of America. It is not covered by the CC-BY 4.0 license, and further reproduction of this figure would need permission from the copyright holder.]

As evident from the diagram in Author response image 4, all of the transgenes inserted on the 3rd chromosome that were tested were able to participate in long-distance (>Mbs) regulatory interactions. On the other hand, not all possible pairwise interactions are observed. This would suggest that potential interactions depend upon the large scale (Mb) 3D folding of the 3rd chromosome.

When the scs boundary (Zw5-dependent, Gaszner et al. 1999) was added to the transgene to give sMws’, it further enhanced the ability of distant transgenes to find each other and pair. All eight of the sMws’ inserts that were tested were able to interact with at least one other sMws’ insert on a different chromosome and silence mini-white. Vazquez et al. (2006) subsequently tagged the sMws’ transgene with LacO sequences (ps0Mws’) and visualized pairing interactions in imaginal discs. Trans-heterozygous combinations on the same chromosome were found paired in 94-99% of the disc nuclei, while a trans-heterozygous combination on different chromosomes was found paired in 96% of the nuclei (Author response image 6). Vazquez et al. (2006) also examined a combination of four transgenes inserted on the same chromosome (two at the same insertion site, and two at different insertion sites). In this case, all four transgenes were clustered together in 94% of the nuclei (Author response image 6). Their studies also suggest that the distant transgenes remain paired for at least several hours. A similar experiment was done by Li et al. (2011), except that the transgene contained only a single boundary, Mcp or Fab-7. While pairing was still observed in trans-heterozygotes, the frequency was reduced without scs and scs’.

Author response image 6.

Author response image 6.

[Author response image 6 is reproduced from Table 3 from Vazquez et al., 2006.]

It is worth pointing out that there is no plausible mechanism in which cohesin could extrude a loop through hundreds of intervening TADs, across the centromere (ff#13.101 with w#11.102: Author response image 4; w#14.29 with w#11.02: Author response images 4 and 5) and come to a halt when it “encounters” Mcp-containing transgenes on different homologs. The same is true for Mcp-dependent pairing interactions in cis (Fig. 7 in Muller et al. 1999) or Mcp-dependent pairing interactions between transgenes inserted on different chromosomes (Line 8 in Author Response Figure 6, Fig. 8 in Muller et al. 1999).

These are not the only boundaries that can engage in long-distance pairing. Mohana et al. (2023) identified nearly 60 meta-loops, many of which appear to be formed by the pairing of TAD boundary elements. Two examples (at 200 bp resolution from 12-16 hr embryos) are shown in Author response image 7.

Author response image 7. Metaloops on the 2nd and 3rd chromosomes: circle-loops and multiple stem-loops.

Author response image 7.

One of these meta-loops (panel A) is generated by the pairing of two TAD boundaries on the 2nd chromosome. The first boundary, blue (indicated by blue arrow), is located at ~2,006,500 bp, between a small TAD containing the Nplp4 and CG15353 genes and a larger TAD containing 3 genes, CG33543, Obp22a, and Npc2a. Nplp4 encodes a neuropeptide. The functions of CG15354 and CG33543 are unknown. Obp22a encodes an odorant binding protein, while Npc2a encodes the Niemann-Pick type C-2a protein that is involved sterol homeostasis. The other boundary (purple: indicated by purple arrow) is located between two TADs 2.8 Mb away at 4,794,250 bp. The upstream TAD contains the fipi gene (CG15630) which has neuronal functions in male courtship, while the downstream TAD contains CG3294, which is thought to be a spliceosome component, and schlaff (slf), which encodes a chitin binding protein. As illustrated in the accompanying diagram, the blue boundary pairs with the purple boundary in a head-to-head orientation, generating a ~2.8 Mb loop with a circle-loop topology. As a result of this pairing, the multi-gene (CG33543, Obp22a, and Npc2a) TAD upstream of the blue boundary interacts with the CG15630 TAD upstream of the purple boundary. Conversely the small Nplp4:CG15353 TAD downstream of the blue boundary interacts with the CG3294:slf TAD downstream of the purple boundary. Even if one imagined that the cohesin bolo tie clip was somehow able to extrude 2.8 Mb of chromatin and then know to stop when it encountered the blue and purple boundaries, it would’ve generated a stem-loop, not a circle-loop.

The second meta-loop (panel B) is more complicated, as it involves pairing interactions between four boundary elements. The blue boundary (blue arrow), located ~4,801,800 bp (3L), separates a large TAD containing the RhoGEF64C gene from a small TAD containing CG7509, which encodes a predicted subunit of an extracellular carboxypeptidase. As can be seen in the MicroC contact profile and the accompanying diagram, the blue boundary pairs with the purple boundary (purple arrow), which is located at ~7,013, 500 (3L), just upstream of the 2nd internal promoter (indicated by black arrowhead) of the Mp (Multiplexin) gene. This pairing interaction is head-to-tail and generates a large stem-loop that spans ~2.2 Mb. The stem-loop brings sequences upstream of the blue boundary and downstream of the purple boundary into contact (like the strings below a bolo tie clip), just as was observed in the boundary bypass experiments of Muravyova et al. (2001) and Kyrchanova et al. (2008). The physical interactions result in a box of contacts (right top) between sequences in the large RhoGEF64C TAD and sequences in a large TAD that contains an internal Mp promoter. The second pairing interaction is between the brown boundary (brown arrow) and the green boundary (green arrow). The brown boundary is located at ~4 805,600 bp (3L), and separates the TAD containing CG7590 from a large TAD containing CG1808 (predicted to encode an oxidoreductase) and the Dhc64C (Dynein heavy chain 64C) gene. The green boundary is located at ~6,995,500 bp (3L), and it separates a TAD containing CG32388 and the biniou (bin) transcription factor from a TAD that contains the most distal promoter of the Mp gene (blue arrowhead). As indicated in the diagram, the brown and green boundaries pair with each other head-to-tail, and this generates a small internal loop (and the final configuration would resemble a bolo tie with two tie clips). This small internal loop brings the CG7590 TAD into contact with the TAD that extends from the distal Mp promoter to the 2nd internal Mp promoter. The resulting contact profile is a rectangular box with diagonal endpoints corresponding to the paired blue:purple and brown:green boundaries. The pairing of the brown:green boundaries also brings the TADs immediately downstream of the brown boundary and upstream of the green boundary into contact with each other, and this gives a rectangular box of interactions between the Dhc64C TAD and sequences in the bin/CG3238 TAD. This box is located on the lower left side of the contact map.

Since the bin and Mp meta-loops in Author response image 7B are stem-loops, they could have been generated by “sequential” cohesin loop extrusion events. Besides the fact that cohesin extrusion of 2 Mb of chromatin and breaking through multiple intervening TAD boundaries challenges the imagination, there is no mechanism in the cohesion loop extrusion/CTCF roadblock model to explain why cohesion complex 1 would come to a halt at the purple boundary on one side and the blue boundary on the other, while cohesin complex 2 would instead stop when it hits the brown and green boundaries. This highlights another problem with the cohesin loop extrusion/CTCF roadblock model, namely that the roadblocks are functionally autonomous: they have an intrinsic ability to block cohesin that is entirely independent of the intrinsic ability of other roadblocks in the neighborhood. As a result, there is no mechanism for generating specificity in loop formation. By contrast, boundary pairing interactions are by definition non-autonomous and depend on the ability of individual boundaries to pair with other boundaries: specificity is built into the model.

The mechanism for pairing, and accordingly the basis for partner preferences/specificity, are reasonably well understood. Probably the most common mechanism in flies is based on shared binding sites for architectural proteins that can form dimers or multimers (Bonchuk et al. 2021; Fedotova et al. 2017). Flies have a large family of polydactyl zinc finger DNA binding proteins, and as noted above, many of these form dimers or multimers, and also function as TAD boundary proteins. This pairing principle was first discovered by Kyrchanova et al. (2008). This paper also showed that orientation-dependent pairing interactions is a common feature of endogenous fly boundaries. Another mechanism for pairing is specific protein:protein interactions between different DNA binding factors (Blanton et al. 2003). Yet a third mechanism would be proteins that bridge different DNA binding proteins together. The boundaries that use these different mechanisms (BX-C boundaries, scs, scs’) depend upon the same sorts of proteins that are used by homie and nhomie. Likewise, this same set of factors reappears, in one combination or another, in most other TAD boundaries. As for the orientation of pairing interactions, this is most likely determined by the order of binding sites for chromosome architectural proteins in the partner boundaries.

…. and many TADs lack focal 3D interactions between their boundaries.

(1.3) The evidence that flies differ from mammals in that they “lack” focal 3D interactions is not compelling. One of the problems with drawing this distinction is that almost all of the “focal 3D interactions” seen mammalian Hi-C experiments are a consequence of binning large DNA segments in low resolution restriction enzyme-dependent experiments. This is even true in the two “high” resolution MicroC experiments that have been published (Hsieh et al. 2020; Krietenstein et al. 2020). As illustrated above in Author response image 1, most of the “focal 3D interactions” (the dots at the apex of TAD triangles) seen with large bin sizes (1 kb and greater) disappear when the bin size is 200 bp, and TADs rather than TAD neighborhoods are being visualized.

As described in point (1.1) above, in the MicroC protocol, fixed chromatin is first digested to mononucleosomes by extensive MNase digestion, processed/biotinylated, and ligated to give dinucleosome-length fragments, which are then sequenced. Regions of chromatin that are nucleosome free (promoters, enhancers, silencers, boundary elements) will typically be reduced to oligonucleotides in this procedure and will not be recovered when dinucleosome-length fragments are sequenced. The loss of sequences from typical paired boundary elements is illustrated by the Lar (Leukocyte-antigen-related-like) meta-loop shown in Author response image 8 (at 200 bp resolution). Panels A and B show the contact profiles generated when the blue boundary (which separates two TADs that span the Lar transcription unit interacts with the purple boundary (which separates two TADs in a gene poor region ~620 kb away)). The blue and purple boundaries pair with each other head-to-head, and this pairing orientation generates yet another circle-loop. In the circle-loop topology, sequences in the TADs upstream of both boundaries come into contact with each other, and this gives the small dark rectangular box to the upper left of the paired boundaries (Author response image 8A). (Note that this small box corresponds to the two small TADs upstream of the blue and purple boundaries, respectively. See panel B.) Sequences in the TADs downstream of the two boundaries also come into contact with each other, and this gives the large box to the lower right of the paired boundaries. While this meta-loop is clearly generated by pairing interactions between the blue and purple boundaries, the interacting sequences are degraded in the MicroC protocol, and sequences corresponding to the blue and purple boundaries aren’t recovered. This can be seen in panel B (red arrow and red arrowheads). When a different Hi-C procedure is used (dHS-C) that captures nucleosome-free regions of chromatin that are physically linked to each other (Author response image 8C, D), the sequences in the interacting blue and purple boundaries are recovered and generate a prominent “dot” at their physical intersection (blue arrow in panel D).

Author response image 8. Lar metaloop.

Author response image 8.

(A, B) MicroC. (C, D) dHS-C.

While sequences corresponding to the blue and purple boundaries are lost in the MicroC procedure, there is at least one class of element that engages in physical pairing interactions whose sequences are (comparatively) resistant to MNase digestion. This class of elements includes many PREs (Kyrchanova et al. 2018; unpublished data), the boundary bypass elements in the Abd-B region of the BX-C (Kyrchanova et al. 2023; Kyrchanova et al. 2019a; Kyrchanova et al. 2019b; Postika et al. 2018), and “tethering” elements (Batut et al. 2022; Li et al. 2023). In all of the cases tested, these elements are bound in nuclear extracts by a large (>1000 kD) GAGA factor-containing multiprotein complex called LBC. LBC also binds to the hsp70 and eve promoters (unpublished data). Indirect end-labeling experiments (Galloni et al. 1993; Samal et al. 1981; Udvardy and Schedl 1984) indicate that the LBC protects a ~120-180 bp DNA segment from MNase digestion. It is likely that this is the reason why LBC-bound sequences can be recovered in MicroC experiments as dots when they are physically linked to each other. One such example (based on the ChIP signatures of the paired elements) is indicated by the green arrow in panel B and D of Author response image 8. Note that there are no dots corresponding to these two LBC elements within either of the TADs immediately downstream of the blue and purple boundaries. Instead the sequences corresponding to the two LBC elements are only recovered when the two elements pair with each other over a distance of ~620 kb. The fact that these two elements pair with each other is consistent with other findings which indicate that, like classical boundaries, LBC elements exhibit partner preferences. In fact, LBC elements can sometimes function as TAD boundaries. For example, the Fab-7 boundary has two LBC elements, and full Fab-7 boundary function can be reconstituted with just these two elements (Kyrchanova et al. 2018).

Reviewer #2 (Public Review):

"Chromatin Structure II: Stem-loops and circle-loops" by Ke*, Fujioka*, Schedl, and Jaynes reports a set of experiments and subsequent analyses focusing on the role of Drosophila boundary elements in shaping 3D genome structure and regulating gene expression. The authors primarily focus on the region of the fly genome containing the even skipped (eve) gene; eve is expressed in a canonical spatial pattern in fly embryos and its locus is flanked by the well-characterized neighbor of homie (nhomie) and homie boundary elements. The main focus of investigation is the orientation dependence of these boundary elements, which had been observed previously using reporter assays. In this study, the authors use Crispr/Cas9 editing followed by recombination-mediated cassette exchange to create a series of recombinant fly lines in which the nhomie boundary element is either replaced with exongenous sequence from phage 𝝀, an inversion of nhomie, or a copy of homie that has the same orientation as the endogenous homie sequence. The nhomie sequence is also regenerated in its native orientation to control for effects introduced by the transgenesis process.

The authors then perform high-resolution Micro-C to analyze 3D structure and couple this with fluorescent and colorimetric RNA in situ hybridization experiments to measure the expression of eve and nearby genes during different stages of fly development. The major findings of these experiments are that total loss of boundary sequence (replacement with 𝝀 DNA) results in major 3D structure changes and the most prominent observed gene changes, while inversion of the nhomie boundary or replacement with homie resulted in more modest effects in terms of 3D structure and gene expression changes and a distinct pattern of gene expression change from the 𝝀 DNA replacement. As the samples in which the nhomie boundary is inverted or replaced with homie have similar Micro-C profiles at the eve locus and show similar patterns of a spurious gene activation relative to the control, the observed effects appear to be driven by the relative orientation of the nhomie and homie boundary elements to one another.

Collectively, the findings reported in the manuscript are of broad interest to the 3D genome field. Although extensive work has gone into characterizing the patterns of 3D genome organization in a whole host of species, the underlying mechanisms that structure genomes and their functional consequences are still poorly understood. The perhaps best understood system, mechanistically, is the coordinated action of CTCF with the cohesin complex, which in vertebrates appears to shape 3D contact maps through a loop extrusion-pausing mechanism that relies on orientation-dependent sequence elements found at the boundaries of interacting chromatin loops.

(2.1) The notion that the mammalian genome is shaped in 3D by the coordinate action of cohesin and CTCF has achieved the status of dogma in the field of chromosome structure in vertebrates. However, as we have pointed out in (1.1), the evidence supporting this dogma is far from convincing. To begin with, it is based on low resolution Hi-C experiments that rely on large bin sizes to visualize so-called “TADs.” In fact, the notion that cohesin and CTCF are responsible on their own for shaping the mammalian 3D genome appears to be a result of mistaking a series of forests for the actual trees that populate each of the forests.

As illustrated in Author response image 1 above, the “TADs” that are visualized in these low resolution data sets are not TADs at all, but rather TAD neighborhoods consisting of several dozen or more individual TADs. Moreover, the “interesting” features that are evident at low resolution (>1 kb), the dots and stripes, largely disappear at resolutions appropriate for visualizing individual TADs (~200 bp).

In Author response image 2, we presented data from one of the key experiments in Goel et al. (2023). In their experiment, the authors used RCMC to generate high resolution MicroC contact maps before and after RAD21 depletion. Contrary to dogma, RAD21 depletion has no effect on TADs in a chromosome 5 DNA segment spanning the Ppm1g gene, when TADs are visualized at 200 bp resolution (as in Author response image 2) or at 250 bp resolution (as in Goel et al. 2023). These TADs look very much like the TADs we observe in the Drosophila genome, in particular, in the Abd-B region of the BX-C that is thought to be assembled into a series of circle-loops (see Fig. 2B).

While Goel et al. (2023) observed no effect of RAD21 depletion on TADs, they found that loss of RAD21 does have some impact on the frequency of longer-distance (but not short-distance) contacts in TAD neighborhoods when their RCMC data set is visualized using bin sizes of 5 kb and 1 kb. This is shown using data from their paper, imaged using a bin size of 1 kb, in Author response image 3. The significance of this finding is, however, uncertain. It could mean that the 3D organization of large TAD neighborhoods have a special requirement for cohesin activity. On the other hand, since cohesin functions to hold sister chromosomes together after replication until they separate during mitosis (and might also participate in mitotic condensation), it is also possible that the loss of long-range contacts in large TAD neighborhoods when RAD21 is depleted is simply a reflection of this particular activity. Further studies will be required to address these possibilities.

As for CTCF: a careful inspection of the ChIP data in Author response image 2 indicates that CTCF is not found at each and every TAD boundary. In fact, the notion that CTCF is the be-all and end-all of TAD boundaries in mammals is truly hard to fathom. For one, the demands for specificity in TAD formation (and in regulatory interactions) are likely much greater than those in flies, and specificity can’t be generated by a single DNA binding protein. For another, several dozen chromosomal architectural proteins have already been identified in flies. This means that (unlike what is thought to be true in mammals) it is possible to use a combinatorial mechanism to generate specificity in, for example, the long-distance interactions in Author response images 7 and 8. As noted in (2.1) above, many of the known chromosomal architectural proteins in flies are polydactyl zinc finger proteins (just like CTCF). There are some 200 different polydactyl zinc finger proteins in flies, and the function of only a hand full of these is known at present. However, it seems likely that a reasonable fraction of this class of DNA binding proteins will ultimately turn out to have an architectural function of some type (Bonchuk et al. 2021; Fedotova et al. 2017). The number of different polydactyl zinc finger protein genes in mammals is nearly 3 times that of flies. It is really possible that of these, only CTCF is involved in shaping the 3D structure of the mammalian genome?

Despite having a CTCF paralog and cohesin, the Drosophila genome does not appear to be structure by loop extrusion-pausing. The identification of orientation-dependent elements with pronounced structural effects on genome folding thus may shed light on alternative mechanisms used to regulated genome structure, which in turn may yield insights into the significance of particular folding patterns.

(2.2) Here we would like to draw the reviewer’s and reader’s attention to Author response image 7, which shows that orientation-dependent pairing interactions have a significant impact on physical interactions between different sequences. We would also refer the reader to two other publications. One of these is Kyrchanova et al. (2008), which was the first to demonstrate that orientation of pairing interactions matters. The second is Fujioka et al. (2016), which describes experiments indicating that nhomie and homie pair with each other head-to-tail and with themselves head-to-head.

On the whole, this study is comprehensive and represents a useful contribution to the 3D genome field. The transgenic lines and Micro-C datasets generated in the course of the work will be valuable resources for the research community. Moreover, the manuscript, while dense in places, is generally clearly written and comprehensive in its description of the work. However, I have a number of comments and critiques of the manuscript, mainly centering on the framing of the experiments and presentation of the Micro-C results and on manner in which the data are analyzed and reported. They are as follows:

Major Points:

(1) The authors motivate much of the introduction and results with hypothetical "stem loop" and "circle loop" models of chromosome confirmation, which they argue are reflected in the Micro-C data and help to explain the observed ISH patterns. While such structures may possibly form, the support for these specific models vs. the many alternatives is not in any way justified. For instance, no consideration is given to important biophysical properties such as persistence length, packing/scaling, and conformational entropy. As the biophysical properties of chromatin are a very trafficked topic both in terms of experimentation and computational modeling and generally considered in the analysis of chromosome conformation data, the study would be strengthened by acknowledgement of this body of work and more direct integration of its findings.

(2.3) The reviewer is not correct in claiming that “stem-loops” and “circle-loops” are “hypothetical.” There is ample evidence that both types of loops are present in eukaryotic genomes, and that loop conformation has significant readouts in terms of not only the physical properties of TADs but also their functional properties. Here we would draw the reviewer’s attention to Author response images 7 and 8 for examples of loops formed by the orientation-dependent pairing of yet other TAD boundary elements. As evident from the MicroC data in these figures, circle-loops and stem-loops have readily distinguishable contact patterns. The experiments in Fujioka et al. (2016) demonstrate that homie and nhomie pair with each other head-to-tail, while they pair with themselves head-to-head. The accompany paper (Bing et al. 2024) also provides evidence that loop topology is reflected both in the pattern of activation of reporters and in the MicroC contact profiles. We would also mention again Kyrchanova et al. (2008), who were the first to report orientation-dependent pairing of endogenous fly boundaries.

At this juncture, it would premature to try to incorporate computational modeling of chromosome conformation in our studies. The reason is that the experimental foundations that would be essential for building accurate models are lacking. As should be evident from Author response images 1-3 above, studies on mammalian chromosomes are simply not of high enough resolution to draw firm conclusions about chromosome conformation: in most studies only the forests are visible. While the situation is better in flies, there are still too many unknown. As just one example, it would be important to know the orientation of the boundary pairing interactions that generate each TAD. While it is possible to infer loop topology from how TADs interact with their neighbors (a plume versus clouds), a conclusive identification of stem- and circle-loops will require a method to unambiguously determine whether a TAD boundary pairs with its neighbor head-to-head or head-to-tail.

(2) Similar to Point 1, while there is a fair amount of discussion of how the observed results are or are not consistent with loop extrusion, there is no discussion of the biophysical forces that are thought to underly compartmentalization such as block-polymer co-segregation and their potential influence. I found this absence surprising, as it is generally accepted that A/B compartmentalization essentially can explain the contact maps observed in Drosophila and other non-vertebrate eukaryotes (Rowley, ..., Corces 2017; PMID 28826674). The manuscript would be strengthened by consideration of this phenomenon.

(2.4) As the reviewer indicates, an alternative mechanism for generating TADs and for explaining the patterns of TAD:TAD interactions is provided by the A/B compartment model. This model is not consistent with the experiments described in either this manuscript or in the accompanying manuscript (Bing et al. 2024). Nor does it fit with extensive studies on the structural and functional properties of boundary elements in the Abd-B region of the bithorax complex. As the reviewer has suggested, we have included a discussion of the A/B compartment model in the last section of the Discussion.

(3) The contact maps presented in the study represent many cells and distinct cell types. It is clear from single-cell Hi-C and multiplexed FISH experiments that chromosome conformation is highly variable even within populations of the same cell, let alone between cell types, with structures such as TADs being entirely absent at the single cell level and only appearing upon pseudobulking. It is difficult to square these observations with the models of relatively static structures depicted here. The authors should provide commentary on this point.

(2.5) As should be evident from Author response image 1, single-cell Hi-C experiments would not provide useful information about the physical organization of individual TADs, TAD boundaries, or how individual TADs interact with their immediate neighbors. In addition, since they capture only a very small fraction of the possible contacts within and between TADs, we suspect that these single-cell studies aren’t likely to be useful for making solid conclusions about TAD neighborhoods like those shown in Author response image 1 panels A, B, C, and D, or Author response image 3. While it might be possible to discern relatively stable contacts between pairs of insulators in single cells with the right experimental protocol, the stabilities/dynamics of these interactions may be better judged by the length of time that physical interactions are seen to persist in live imaging studies such as Chen et al. (2018), Vazquez et al. (2006) and Li et al. (2011).

The in situ FISH data we’ve seen also seems problematic in that probe hybridization results in a significant de-condensation of chromatin. For two probe sets complementary to adjacent ~1.2 kb DNA sequences, the measured center-to-center distance that we’ve seen was ~110 nM. This is about 1/3 the length that is expected for a 1.2 kb naked DNA fragment, and about 1.7 times larger than that expected for a beads-on-a-string nucleosome array (~60 nM). However, chromatin is thought to be compacted into a 30 nM fiber, which is estimated to reduce the length of DNA by at least another ~6 fold. If this estimate is correct, FISH hybridization would appear to result in a ~10 fold decompaction of chromatin. A decompaction of this magnitude would necessarily be followed by a significant distortion in the actual conformation of chromatin loops.

(4) The analysis of the Micro-C data appears to be largely qualitative. Key information about the number of reads sequenced, reaps mapped, and data quality are not presented. No quantitative framework for identifying features such as the "plumes" is described. The study and its findings would be strengthened by a more rigorous analysis of these rich datasets, including the use of systematic thresholds for calling patterns of organization in the data.

Additional information on the number of reads and data quality have been included in the Methods section.

(5) Related to Point 4, the lack of quantitative details about the Micro-C data make it difficult to evaluate if the changes observed are due to biological or technical factors. It is essential that the authors provide quantitative means of controlling for factors like sampling depth, normalization, and data quality between the samples.

The reviewer suggests that biological and/or technical differences between the four samples could account for the observed changes in the MicroC patterns for the eve TAD and its neighbors. If this were the case, then similar changes in MicroC patterns should be observed elsewhere in the genome. Since much of the genome is analyzed in these MicroC experiments, there is an abundance of internal controls for each experimental manipulation of the nhomie boundary. For two of the nhomie replacements, nhomie reverse and homie forward, the plume above the eve volcano triangle is replaced by clouds surrounding the eve volcano triangle. If these changes in the eve MicroC contact patterns are due to significant technical (or biological) factors, we should observe precisely the same sorts of changes in TADs elsewhere in the genome that are volcano triangles with plumes.

Author response image 9 shows the MicroC contact pattern for several genes in the Antennapedia complex. The deformed gene is included in a TAD which, like eve, is a volcano triangle topped by a plume. A comparison of the deformed MicroC contact patterns for nhomie forward (panel B) with the MicroC patterns for nhomie reverse (panel C) and homie forward (panel D) indicates that while there are clearly technical differences between the samples, these differences do not result in the conversion of the deformed plume into clouds as is observed for the eve TAD. The MicroC patterns elsewhere in Antennapedia complex are also very similar in all four samples. Likewise, comparisons of regions elsewhere in the fly genome indicate that the basic contact patterns are similar in all four samples. So while there are technical differences which are reflected in the relative pixel density in the TAD triangles and the LDC domains, these differences do not result in converting plumes into clouds nor do the alter the basic patterns of TAD triangles and LDC domains. As for biological differences—the embryos in each sample are at roughly the same developmental stage and were collected and processed using the same procedures. Thus, the biological factors that could reasonably be expected to impact the organization of specific TADs (e.g., cell-type specific differences) are not going to impact the patterns we see in our experiments.

Author response image 9.

Author response image 9.

(6) The ISH effects reported are modest, especially in the case of the HCR. The details provided for how the imaging data were acquired and analyzed are minimal, which makes evaluating them challenging. It would strengthen the study to provide much more detail about the acquisition and analysis and to include depiction of intermediates in the analysis process, e.g. the showing segmentation of stripes.

The imaging analysis presented in Fig. 5 is just standard confocal microscopy. Individual embryos were visualized and scored. An embryo in which stripes could be readily detected was scored as ‘positive’ while an embryo in which stripes couldn’t be detected was scored as ‘negative.’

Recommendations for the authors:

Editor comments:

It was noted that the Jaynes lab previously published extensive genetic evidence to support the stem loop and circle loop models of Homie-Nhomie interactions (Fujioka 2016 Plos Genetics) that were more convincing than the Micro-C data presented here in proof of their prior model. Maybe the authors could more clearly summarize their prior genetic results to further try to convince the reader about the validity of their model.

Reviewer #1 (Recommendations For The Authors):

Below, I list specific comments to further improve the manuscript for publication. Most importantly, I recommend the authors tone down their proposal that boundary pairing is a universal TAD forming mechanism.

(1) The title is cryptic.

The title has been changed.

(2) The second sentence in the abstract is an overstatement: "In flies, TADs are formed by physical interactions between neighboring boundaries". Hi-C and Micro-C studies have not provided evidence that most TADs in Drosophila show focal interactions between their bracketing boundaries. The authors rely too strongly on prior studies that used artificial reporter transgenes to show that multimerized insulator protein binding sites or some endogenous fly boundaries can mediate boundary bypass, as evidence that endogenous boundaries pair.

Please see responses (1.1) and (1.3) and Author response images 1 and 7. Note that using dHS-C, most TADs that we’ve looked at so far are topped by a “dot” at their apex.

(3) Line 64: the references do not cite the stated "studies dating back to the '90's'".

The papers cited for that sentence are reviews which discussed the earlier findings. The relevant publications are cited at the appropriate places in the same paragraph.

(4) Line 93: "On the other hand, while boundaries have partner preferences, they are also promiscuous in their ability to establish functional interactions with other boundaries." It was unclear what is meant here.

Boundaries that (a) share binding sites for proteins that multimerize, (b) have binding sites for proteins that interact with each other, or (c) have binding sites for proteins that can be bridged by a third protein can potentially pair with each other. However, while these mechanisms enable promiscuous pairing interactions, they will also generate partner preferences (through a greater number of a, b, and/or c).

(5) It could be interesting to discuss the fact that it remains unclear whether Nhomie and Homie pair in cis or in trans, given that homologous chromosomes are paired in Drosophila.

The studies in Fujioka et al. (2016) show that nhomie and homie can pair both in cis and in trans. Given the results described in (1.2), we imagine that they are paired both in cis and in trans in our experiments.

(6) Line 321: Could the authors further explain why they think that "the nhomie reverse circle-loop also differs from the nhomie deletion (λ DNA) in that there is not such an obvious preference for which eve enhancers activate expression"?

The likely explanation is that the topology/folding of the altered TADs impacts the probability of interactions between the various eve enhancers and the promoters of the flanking genes. We have added a phrase to that sentence to make this more clear.

(7) The manuscript would benefit from shortening the long Discussion by avoiding repeating points described previously in the Results.

(8) Line 495: "If, as seems likely, a significant fraction of the TADs genome-wide are circle loops, this would effectively exclude cohesin-based loop extrusion as a general mechanism for TAD formation in flies". The evidence provided in this manuscript appears insufficient to discard ample evidence from multiple laboratories that TADs form by compartmentalization or loop extrusion. Multiple laboratories have, for example, demonstrated that cohesin depletion disrupts a large fraction of mammalian TADs.

Points made here and in #9 have been responded to in (1.1), (2.1), and (2.4) above. We would suggest that the evidence for loop extrusion falls short of compelling (as it is based on the analysis of TAD neighborhoods, not TADs—that is forests, not trees) and, given the results reported in Goel et al. (in particular Fig. 4 and Sup. Fig. 8), is clearly suspect. This is not to mention the fact that cohesin loop extrusion can’t generate circle-loop TADs, yet circle-loops clearly exist. Likewise, as discussed in (2.4), it is not clear to us that the shared chromatin states, polymer co-segregation and co-repulsion, account for the compartmental patchwork patterns of TAD:TAD interactions. The results from the experimental manipulations in this paper and the accompanying paper, together with studies by others, e.g., Kyrchanova et al. (2008) and Mohana et al. (2023), would also seem to be at odds with the model for compartments as currently formulated.

The unique properties of Nhomie and Homie, namely the remarkable specificity with which they physically pair over large distances (Fujioka et al. 2016) may rather suggest that boundary pairing is a phenomenon restricted to special loci. Moreover, it has not yet been demonstrated that Nhomie or Homie are also able to pair with the TAD boundaries on their left or right, respectively.

Points made here were discussed in detail in (1.2). As described in detail in (1.2), it is not the case that nhomie and homie are “unique” or “special.” Other fly boundaries can do the same things. As for whether nhomie and homie pair with their neighbors: we haven’t done transgene experiments (e.g., testing by transvection or boundary bypass). Likewise, in MicroC experiments there are no obvious dots at the apex of the neighboring TADs that would correspond to nhomie pairing with the neighboring boundary to the left and homie pairing with the neighboring boundary to the right. However, this is to be expected. As we discussed in (1.3) above, only MNase-resistant elements will generate dots in standard MicroC experiments. On the other hand, when boundary:boundary interactions are analyzed by dHS-C (c.f., Author response image 8), there are dots at the apex of both neighboring TADs.

(9) The comment in point 8 also applies to the concluding 2 sentences (lines 519-524) of the Discussion.

See the response to #8 above. Otherwise, the concluding sentences are completely accurate. Validation of the cohesin loop-extrusion/CTCF roadblock model will required demonstrating (a) that all TADs are either stem-loops or unanchored loops, and (b) that TAD endpoints are always marked by CTCF.

The likely presence of circle-loops and the evidence of TAD boundaries that don’t have CTCF (c.f., Author response image 2 from Goel et al. 2023) already suggest that this model can’t (either fully or perhaps not even primarily) account for TAD formation in mammals.

(10) Figs. 3 and 6: It would be helpful to add the WT screenshot in the same figure, for direct comparison.

We think it is easy enough to scroll between figures, especially since nhomie forward looks just like WT.

(11) Fig. 6: It would be helpful to show a cartoon view of a circle loop to the right of the Micro-C screenshot, as was done in Fig. 3.

Good idea. This was added to the figure.

(12) Fig. 5: It would be helpful to standardize the labelling of the different genotypes throughout the figures and panels ("inverted" versus "reverse" versus an arrow indicating the direction).

This was fixed.

Reviewer #2 (Recommendations For The Authors):

Minor Points:

(1) The Micro-C data does not appear to be deposited in an appropriate repository. It would be beneficial to the community to make these data available in this way.

This has been done.

(2) Readers not familiar with Drosophila development would benefit from a gentle introduction to the stages analyzed and some brief discussion on how the phenomenon of somatic homolog pairing might influence the study, if at all.

We included a rough description of the stages that were analyzed for both the in situs and MicroC. We thought that a full description of what is going on at each of the stages wasn’t necessary, as the process of development is not a focus of this manuscript. In other studies, we’ve found that there are only minor differences in MicroC patterns between the blastoderm stage and stage 12-16 embryos. While these minor differences are clearly interesting, we didn’t discuss them in the text. In all of our experiments, chromosomes are likely to be paired. In NC14 embryos (the stage for visualizing eve stripes and the MicroC contact profiles in Fig. 2) replication of euchromatic sequences is thought to be quite rapid. While homolog pairing is incomplete at this stage, sister chromosomes are paired. In stage 12-16 embryos, homologs will be paired, and if the cells are arrested in G2, then sister chromosome will also be paired. So, in all of the experiments, chromosomes (sisters and/or homologs) are paired. However, since we don’t have examples of unpaired chromosomes, our experiments don’t provide any info on how chromosome pairing might impact MicroC/expression patterns.

(3) "P > 0.01" appears several times. I believe the authors mean to report "P < 0.01".

This was fixed.

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Associated Data

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

    Data Citations

    1. Ke W, Fujioka M, Schedl P, Jaynes J. 2024. Chromosome Structure II: Stem-loops and circle-loops. NCBI Gene Expression Omnibus. GSE263270
    2. Ke W. 2024. Stem-loop and circle-loop TADs generated by directional pairing of boundary elements have distinct physical and regulatory properties. Open Science Framework. 6pybm/ [DOI] [PMC free article] [PubMed]
    3. Bing X, Batut P, Levine M. 2022. Genome organization controls transcriptional dynamics during development. NCBI Gene Expression Omnibus. GSE171396 [DOI] [PMC free article] [PubMed]
    4. Bing X, Levo M, Raimundo J, Levine M. 2022. Transcriptional coupling of distant regulatory genes in living embryos. NCBI Gene Expression Omnibus. GSE173518 [DOI] [PMC free article] [PubMed]

    Supplementary Materials

    MDAR checklist

    Data Availability Statement

    Sequence data are available at GEO GSE263270. Confocal images are available on Open Science Framework at https://doi.org/10.17605/OSF.IO/6PYBM.

    The following datasets were generated:

    Ke W, Fujioka M, Schedl P, Jaynes J. 2024. Chromosome Structure II: Stem-loops and circle-loops. NCBI Gene Expression Omnibus. GSE263270

    Ke W. 2024. Stem-loop and circle-loop TADs generated by directional pairing of boundary elements have distinct physical and regulatory properties. Open Science Framework. 6pybm/

    The following previously published datasets were used:

    Bing X, Batut P, Levine M. 2022. Genome organization controls transcriptional dynamics during development. NCBI Gene Expression Omnibus. GSE171396

    Bing X, Levo M, Raimundo J, Levine M. 2022. Transcriptional coupling of distant regulatory genes in living embryos. NCBI Gene Expression Omnibus. GSE173518


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