Introduction

The mammalian genomic DNA that is roughly 2 meters long in a cell is folded extensively in order to fit the size of the nucleus with a diameter of ~5–10 μm.1 Microscopic and 3C-based approaches reveal a hierarchical organization of the genome.2,3,4,5 At the megabase scale, chromatin is subdivided into two spatially segregated compartments, arbitrarily labeled as A and B, with distinct transcriptional activity and histone modification as well as other features such as CpG frequency and DNA replication timing.6,7,8,9,10 The euchromatic A compartment adopts a central position, whereas the heterochromatic B compartment moves towards the nuclear periphery and nucleolar regions.11 This nuclear organization appears to be conserved from ciliates to humans and has been maintained in eukaryotes over 500 million years of evolution.12 Within compartments at the kilobase-to-megabase scale, chromatin is organized in topologically associated domains (TADs), which serve as functional platforms for physical interactions between co-regulated genes and regulatory elements.13 At a finer scale, TADs are divided into smaller loop domains, in which distal regulatory elements such as enhancers come into direct contact with their target genes via chromatin loops.14 Intriguingly, most A/B compartments and TADs are relatively stable in different mouse and human cell types (Supplementary information, Text S1), whereas sub-TAD loops and a small fraction of lineage-specific regions with less pronounced compartment associations tend to be more variable for differential gene expression during cell-fate transition.13,15,16,17,18,19

Evidence suggests that compartments and TADs may be formed by distinct mechanisms. TADs are thought to be formed by active extrusion of chromatin loops by the ring-shaped cohesin complex, which co-localizes with the insulator protein CTCF at the boundaries and anchor regions of contact domains and loops.20,21 Depletion of CTCF disrupted TAD boundaries but failed to impact compartmentalization, whereas cohesion loss made TADs disappear but increased compartmentalization, although both eliminated sub-TAD loop contacts.22,23,24,25,26,27,28,29,30 These results indicate that compartmentalization of mammalian chromosomes emerges independently of proper insulation of TADs. A few mechanisms have been proposed for compartmentalization, such as anchoring heterochromatin to the nuclear lamina,31,32,33,34,35 preferential attraction of chromatin harboring similar histone modifications and regulators,4,36,37,38,39 and hypothetical models involving pairing of homologous sequences mediated by active transcription and phase separation of block copolymers.36,40,41,42,43,44 Although lamin-associated domains (LADs) contribute to a basal chromosome architecture, a large body of work has demonstrated a secondary role for lamina scaffolding in compartmental segregation of heterochromatin and euchromatin.31,32,33,34,35,45,46,47,48,49 In vitro assembled nucleosomal arrays harboring histone H3 lysine 9 di- and tri-methylation (H3K9me2 and H3K9me3) marks undergo phase separation with heterochromatin protein 1 (HP1) and associated proteins to form macromolecule-enriched liquid droplets, reminiscent of heterochromatin.38 However, the role of histone modifications in regulating compartmentalization in vivo remains uncertain. Taken SUV39H H3K9 methyltransferases for example, SUV39H double-null cells still exhibit DAPI-dense heterochromatin foci despite the loss of pericentric H3K9me3 marks;39 and double knockout mice of SUV39H survive at birth with abnormalities.48 A phase-separation model of block copolymers with similar activity appears attractive in explaining compartmental formation.12,34,40,41,42,43,44 However, this hypothesis remains inconclusive, owing to a large void of identification and experimental validation of the molecular drivers that underlie compartmental segregation of euchromatin and heterochromatin.

Repetitive elements comprise more than half of human and mouse genomes.50,51 Once regarded as genomic parasites,52 retrotransposons have been recently implicated in playing active roles in re-wiring the genome and gene expression programs in diverse biological processes.53,54,55,56,57,58,59,60,61 Long and short interspersed nuclear elements (LINEs and SINEs, respectively) are the two predominant subfamilies of retrotransposons in most mammals.62 L1 (also named as LINE1 or LINE-1) is the most abundant subclass of all repeats, making up to 19% and 17% (0.9–1.0 million copies) of the genome in mouse and human, respectively.63 B1 in mouse and its closely related, primate-specific Alu elements in human are the most abundant subclass of SINEs, constituting 3%–11% (0.6–1.3 million copies) of mouse and human genomes.64,65 L1 and B1/Alu have distinct nucleotide compositions and sequence lengths. L1 elements are 6–7 kb long and AT-rich, while Alu elements are ~300 bp long and rich in G and C nucleotides.66 Analysis of metaphase chromosome banding showed roughly inverse distributions of L1 and Alu elements in chromosomal regions with distinct biochemical properties.45,67,68 Initial studies suggested that Alu/B1 elements appear to be enriched in gene-rich, euchromatic A compartments, whereas L1 elements tend to be enriched in gene-poor, heterochromatic B compartments that interact with lamina-associated domains.35,45,47,69,70,71 However, evidence to pinpoint a role for L1 and B1/Alu repeats in organizing the genome has to our knowledge not been reported, albeit fragmented information about their localizations in scattered reports (Supplementary information, Text S1). Systematic map** and visualization of L1 and B1/Alu distributions are still lacking.

We have postulated that the primary DNA sequences, particularly abundant repetitive elements embedded in the genome, may instruct genome folding.61 Here, we report that L1 and B1/Alu repeats tend to cluster with sequences from their own repeat subfamily and form grossly exclusive domains in the nuclear space, which efficiently explains and predicts the compartmental organization revealed by Hi-C. The segregated pattern of L1-rich sequences in the nuclear and nucleolar peripheries and B1/Alu-rich sequences in the nuclear interior is highly conserved across a variety of mouse and human cells, and re-occurs during the cell cycle. In addition, de novo establishment of nuclear segregation of L1- and B1-rich compartments is coincident with the formation of higher-order chromatin structures during early embryogenesis, and appears to be critically regulated by L1 and B1 repeat RNA. Importantly, depletion of L1 RNA in mouse embryonic stem cells (mESCs) significantly weakens spatial contacts of homotypic repeat DNA, disrupts the nuclear localization and segregation of L1- or B1-rich chromosomal sequences, and leads to attenuated compartmentalization of the higher-order chromatin structure. Moreover, we show that recombinant HP1α is able to bind RNA and to phase separate in the presence of RNA or DNA in vitro. Genome-wide co-localization of L1 and HP1α renders these repeat DNA and RNA sequences an advantage in promoting HP1α phase separation in heterochromatin contexts. Altogether, our findings suggest a genetically encoded mechanism by which L1 and B1/Alu repeats organize chromatin macrostructure at the compartmental level, providing an important clue to the conservation and robustness of the higher-order chromatin structure across mouse and human.

Results

L1 and B1/Alu distributions correlate with global compartmentalization in mouse and human

We analyzed the genomic positions of the major repeat subfamilies in mouse and observed positive correlations within L1 or SINE B1 subfamilies, but strong inverse correlations between them (Supplementary information, Fig. S1a). This suggests that L1 and B1 elements tend to be positioned away from each other in the genome, while repeats from the same subfamily tend to be clustered. The non-random positioning of repeat sequences in the genome prompted us to examine their relative distributions in high-order chromatin structures. We first analyzed the published Hi-C data from mESCs.72 Dense L1 and B1 repeats appear to be enriched in distinct compartments across the mouse genome, and within a compartment they are evenly distributed without obvious bias towards the boundary (Fig. 1a, b). L2 repeats show weak enrichments in B1-rich compartments, whereas other types of retrotransposons such as ERV1 and ERVK tend to be randomly distributed (Fig. 1a and Supplementary information, Fig. S1b). The compartments marked by B1 repeats show enrichment of active histone marks (H3K4me3, H3K9ac, H3K27ac, H3K36me3), strong binding of RNA polymerase II (Pol II), and high levels of chromatin accessibility and transcription activity. In contrast, the compartments marked by L1 repeats show signatures of heterochromatin, including enrichment of the repressive H3K9me2 and H3K9me3 marks, and strong binding of heterochromatin proteins such as HP1α and the nuclear corepressor KRAB-associated protein-1 (KAP1 or TRIM28) (Fig. 1a and Supplementary information, Fig. S1b).

Fig. 1: B1- and L1-rich genomic regions homotypically interact, characterize and predict Hi-C compartments.
figure 1

a Heatmaps of the distribution densities of B1, L1, L2, and ERV1 repeats and random genomic regions (panel (i)), DNase I hypersensitive sites (DHS) and ChIP-seq signals of Pol II, H3K4me3, HP1α, and H3K9me3 (panel (ii)), ChIRP-seq signals of Malat1 and L1 RNA (panel (iii)), and RNA-seq (panel (iv)) in mESCs across two adjacent compartments (Cn, Cn+1). All signals in 696 compartments annotated in mESCs were sorted according to the B1 distributions shown in panel (i). b Relative contents of L1 and B1/Alu repeats across the A and B compartments annotated in various cell types in mouse (top) and human (bottom). Random genomic regions serve as the negative control. c Genome browser shots showing conserved domain structures as indicated by heatmaps of the Hi-C contact matrix over a syntenic region in mouse (top) and human (bottom) ESCs. The B1/Alu and L1 repeat densities, the A/B compartments are shown by eigenvalues of the Hi-C contact matrix, and Refseq gene annotations are shown underneath each heatmap. d Heatmap of normalized interaction frequencies at 100-kb resolution on chromosome 17 in mESCs. Genomic distributions and densities of B1 and L1 repeats are shown in the left and bottom tracks. e A zoomed-in view of the interaction matrix of the genomic region from 18 to 60 Mb on mouse chr17 (40-kb resolution). Under the heatmap, we show sequentially genomic distributions and densities of B1 and L1 repeats (in 10-kb bin), log2 ratio of B1 to L1 density, eigenvalues of the Hi-C matrix representing A/B compartments from mouse mESCs, neural progenitor cells (NPC) and neurons, and Pol II ChIP-seq signals and annotated TADs in mESCs. B1-rich regions are arbitrarily labeled as D, F, and H in uppercase. L1-rich regions are labeled as c, e, g, and i in lowercase. Some strong homotypic interactions between compartments rich in the same repeat subfamily (for example, between the B1-rich regions DF, DE and FH, and between the L1-rich regions ce and cg), are highlighted by dotted boxes. f Correlation heatmap showing Pearsonʼs correlation coefficients of the interaction frequencies of any two paired regions in a sub-region on chr17 (500-kb resolution). B1-rich regions are labeled in uppercase as F, H … R, T. L1-rich regions are labeled in lowercase as e, g … q, u. Dotted boxes (in red) and arrows highlight positive correlations of the anchor region F (indicated by *) with other B1-rich genomic regions (horizontal), and of the anchor region g (indicated by *) with other L1-rich genomic regions (vertical). g De novo compartment calling based on L1 and B1 DNA sequences. Panel (i) shows the percentage of L1- or B1-rich compartments overlapped with A or B compartments identified by Hi-C. Panel (ii) shows representative genomic regions with ratio of B1 to L1 in log2 scale and PC1 score of Hi-C interaction matrix.

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We then performed a quantitative sequence analysis of annotated A/B compartments in six distinct mouse and human cell types.20,72 All cells exhibit consistently high levels of SINE repeats (including B1, B2, B4 in mouse and Alu in human) in the A compartments and L1 repeats (including truncated or intact, and evolutionary old or young L1s) in the B compartments (Fig. 1b and Supplementary information, Fig. S1c–e). In contrast, L2 and ERV1 repeats fail to show consistent enrichments across mouse and human. In addition, unsupervised clustering revealed that the genomic positions of A/B compartments are highly similar across six cell types, with an average Spearman correlation coefficient > 0.73 within species and > 0.52 between species (Supplementary information, Fig. S1f). Compared to other subclass repeats, L1 and B1/Alu are most strongly related to the high-order chromatin structures, and their distributions appear to be conserved in homologous regions of the mouse and human genomes (Supplementary information, Fig. S1f). For example, a region in mouse chromosome 2 (chr2: 140–170 Mb) and its syntenic region in human chromosome 20 (chr20: 6–50 Mb) show similar patterns of Hi-C contact probabilities, and gene and repeat compositions and distributions in the corresponding A and B compartments along the DNA sequences (Fig. 1c).

We further analyzed the published datasets of higher-order chromatin interactions in 21 primary human tissues and cell lines.17 On the basis of the PC1 values of a principal components analysis on the Hi-C correlation matrix reported by Schmitt et al.,17 we found that A/B-compartmental associations are highly correlated across all 21 examined samples with correlation coefficients ranging 0.47–0.99 and a median value of 0.79 (Supplementary information, Fig. S2a). The degree of compartmental conservation is highly significant (P < 2.2e-16), as ~80% of the genome shows consistent compartmental labeling in at least 16 samples and ~40% is invariant in all 21 samples, in contrast to 7% and 0% to be expected by chance, respectively (Supplementary information, Fig. S2b, c). Most of compartmental switches that account for 20% of the genome occur in one or few (≤ 5) samples with less pronounced compartmental labeling (gray highlighted regions with low absolute values of PC1 in Supplementary information, Fig. S2d; see also Supplementary information, Text S1). Thus, despite some switching events occurring in individual cells, global compartmentalization is rather stable. Consistently, the genomic regions with conserved A or B compartments across samples exhibit significantly higher levels of Alu or L1 repeats, respectively (Supplementary information, Fig. S2e). Altogether, these results indicate that co-segregation of B1/Alu and L1 repeats with the A and B compartments appears to be stable in different cell types in mouse and human.

Homotypic clustering of L1 and B1 repeats characterizes and predicts compartmental organization

To have a close look at repeat distribution and the higher-order chromatin structure, we took mouse chromosome 17 (chr17, 95 Mb in length) as an example to overlay L1 and B1 features on the Hi-C interaction matrix of mESCs. Interestingly, the plaid pattern of enriched and depleted interaction blocks in the Hi-C map is largely correlated with the compositions and distributions of B1 and L1 along the whole chr17 (Fig. 1d). In a 42-Mb region of chr17, four L1-rich compartments (denoted by c, e, g and i) and three B1-rich compartments (denoted by D, F and H) are alternately positioned along the linear DNA sequence (Fig. 1e). Strong interactions were observed within L1-rich compartments (represented by ce, cg, eg, ei and gi, dotted boxes) or B1-rich compartments (represented by DF, DH and FH, dotted boxes), but not between these two compartments (Fig. 1e). The interaction frequencies between D and F (DF) or between c and e (ce) are much stronger than those of D or F with c or e (cD, De, eF), despite the fact that these regions are closer in the linear sequence. Note that L1- or B1-rich segments often span several adjacent TADs (Fig. 1e and Supplementary information, Fig. S2f), consistent with the findings that TADs are smaller, structural units of compartments.73,74 L1 and B1 compositions within a TAD also exhibit strong anti-correlations (Pearson correlation coefficient <−0.75) across 2200 annotated TADs in mESCs (Supplementary information, Fig. S2g). This observation is consistent with repeat analyses at the genome-wide and compartmental levels (Fig. 1a and Supplementary information, Fig. S1a), illustrating a mutually exclusive distribution of L1 or B1-rich sequences along the genome.

Conversion of Hi-C contact frequencies into Pearson correlation coefficients sharpened our view of the long-range chromatin interactions (Fig. 1f). By visual inspection, we found that the plaid pattern of the Hi-C correlation map precisely matches the distribution and interaction status of L1 and B1. L1-rich or B1-rich regions show strong enrichment of contacts with regions containing the same repeat type (red blocks in Fig. 1f). We refer to these as homotypic contacts. Contacts between regions containing the other repeat type (heterotypic interactions) are strongly depleted (blue blocks in Fig. 1f). For example, in one region of chr17 (35 to 95 Mb), L1-rich segments (from e to u) and B1-rich segments (from F to T) exhibit high frequencies of homotypic contacts (Fig. 1f, highlighted by arrows), but strong depletion of heterotypic contacts. Similarly, homotypic contacts between L1-rich regions or B1-rich regions were also observed between chromosomes, as illustrated by chromosomes 17 and 19 (Supplementary information, Fig. S2h). These results indicate that genomic regions containing B1 or L1 repeats tend to interact with genomic regions containing repeat sequences from similar subfamilies, but not from different subfamilies, regardless of linear proximity, which characterizes the organization at intra- and inter-chromosomal levels.

Next, we sought to predict compartmental organization based on repeat distributions. We used the criterion of log2 ratio of B1 to L1 density [log2(B1/L1)] larger or smaller than 0 for B1-rich or L1-rich compartments, respectively. About 540 B1-rich and 648 L1-rich compartments were identified with a median size of 1.2 Mb across the mouse genome (Supplementary information, Table S1). The numbers and sizes of these B1- and L1-rich compartments called de novo are comparable to those of A and B compartments annotated by Hi-C in mESCs (366 and 364, respectively, with a median size of 1.9 Mb). Importantly, 82% of B1-rich compartments and 77% of L1-rich compartments are overlapped with annotated A or B compartments, respectively (Fig. 1g and Supplementary information, Fig. S3). Only 18% to 23% of compartments show inconsistent labeling between our prediction and Hi-C. We then analyzed genomic features in these ‘falsely’ labeled regions. Intriguingly, L1-rich regions that fall into Hi-C-annotated A compartments (designated as ‘L1.A’) still exhibit a high level of heterochromatic H3K9me3 mark and low levels of chromatin accessibility and gene expression, and contain genes enriched in specialized functions such as responses to pheromone and immunoglobulin and synapse (Supplementary information, Fig. S4a–c). Similarly, B1-rich regions that fall into Hi-C-annotated B compartments (designated as ‘B1.B’) exhibit high levels of chromatin accessibility and gene expression but low H3K9me3 binding (Supplementary information, Fig. S4a–c). In addition, L1.A and B1.B regions exhibit significantly less pronounced PC1 values (close to zero) than those consistent regions (B1.A and L1.B) (Supplementary information, Fig. S4a). Thus, a mere usage of B1 to L1 density ratios successfully re-constructs most of A and B compartments annotated by Hi-C, which suggests that the linear genomic DNA repeats contain the macroscopic structural information. Taken together, homotypic clustering of regions rich in B1 or L1 repeats nicely explains and predicts genome organization at the compartmental level.

Nuclear segregation of L1- and B1/Alu-rich compartments is conserved

High-resolution imaging of L1 and B1 distributions in the conventional nucleus remains lacking, despite initial evidence of their differential localization.45,68 To visualize their positioning in the nuclear space, we performed dual-color DNA fluorescence in situ hybridization (FISH) using fluorescence-tagged oligonucleotide probes that specifically target the consensus sequences of B1 and L1 elements (Fig. 2a). Strikingly, L1 and B1 exhibit distinct yet complementary nuclear localizations in mESCs (Fig. 2b and Supplementary information, Fig. S5a). B1 DNA shows punctate signals in the nuclear interior. In contrast, L1 DNA exhibits highly organized and concentrated signals that line the periphery of the nucleus and nucleolus. Weak L1 signals were also detected in a few areas of the nuclear interior subregions where B1 signals were absent. Both B1 and L1 signals are absent from DAPI-dense regions, which likely represent satellite repeat-enriched chromocenters.75

Fig. 2: DNA FISH reveals the spatial segregation of L1 and B1 compartments.
figure 2

a Schematic illustrations of L1 (panel (i)) and B1 (panel (ii)) RNA targeted by AMO/ASO or DNA FISH probes. b Representative images of L1 (green) and B1 (red) repeats revealed by DNA FISH in mESCs. DNA is labeled by DAPI (blue). All scale bars, 5 μm. c Representative images of DNA immuno-FISH analysis of L1 (green) and B1 (red) DNA repeats, and NCL protein (purple) in mESCs (n = 37), NSC (n = 23), NIH3T3 (n = 18), C2C12 (n = 11) and HeLa cells (n = 15). d Representative images of Oligopaint DNA FISH of individual sites in A or B compartments. Panel (i), three A-compartmental regions (F, H, R) with B1 DNA FISH; panel (ii), two B-compartmental regions (g, q) with L1 DNA FISH. See also Supplementary information, Fig. S5.

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To confirm the L1 localization at the nucleolar periphery in mESCs, we performed DNA immuno-FISH, using an antibody against the nucleolar marker Nucleolin (NCL). Indeed, L1 signals surround and partially overlap with the ring-shaped signals of NCL at the nucleolar periphery (Fig. 2c). The localization of L1 surrounding the nucleus and nucleolus is consistent with sequencing-based analysis of nucleolus- and lamina-associated domains (NADs and LADs), in which L1-rich sequences are sequestered.61,69,76,77 In addition, to further confirm nuclear colocalization of L1-rich sequences in B compartments and B1-rich sequences in A compartments, we performed Oligopaint DNA FISH for five representative loci, each of which ranging from ~100 to 1 Mb was targeted by a set of 500–4500 DNA probes (targeting single-copy sequences) at a density of 200–300 bp per probe. Indeed, three regions (F, H, and R) annotated in the A compartment are colocalized with B1 FISH signals in the nuclear interior (205 out of 217 nuclei), whereas two B compartment-associated regions (g and q) are colocalized with L1 FISH signals in either LAD or NAD (248 out of 251 nuclei) (Fig. 2d and Supplementary information, Fig. S5b, c).

Moreover, to ask whether L1 and B1 localizations might vary with cell type, we analyzed four additional cell lines, including mouse neural stem cells (NSC), fibroblasts (NIH3T3), and myoblasts (C2C12), and human cervical cancer cells (HeLa) (Fig. 2c). Similar to mESCs, all these cells show non-overlap** localizations of B1/Alu in the nuclear interior and L1 at the nuclear and nucleolar peripheries. Thus, consistent with Hi-C results, the segregated staining pattern of B1/Alu and L1 further demonstrates that homotypic clustering of similar repeat sequences in the nuclear space divides the nucleus into distinct territories. This pattern is conserved across different cell types in mouse and human.

Dynamic re-construction of L1 and B1/Alu segregation during the cell cycle

We then asked whether the nuclear segregation of L1- and B1-rich compartments could be re-constructed during mitosis when chromatin structure undergoes dynamic reorganization. DNA FISH analysis of synchronous mESCs showed that L1 and B1 localizations change dramatically at different cell cycle stages (Fig. 3a and Supplementary information, Fig. S6a). S-phase cells show non-overlap** and complementary localizations of L1 and B1 repeats (Figs. 2b and 3a). This is similar to the pattern we observed previously in asynchronous mESCs, more than half of which are in the S phase of the cell cycle (Supplementary information, Fig. S6a). However, L1 and B1 DNA signals are mixed on mitotic chromosomes in metaphase (M phase, including prophase and anaphase), when the nuclear membrane and nucleoli are disassembled. As the cell cycle progresses into the G1 phase, L1 and B1 DNA start to segregate again (Fig. 3a and Supplementary information, Fig. S6b). To quantify the degree of segregation, we defined a FISH-based segregation index as the negative value of Pearson’s correlation coefficient of L1 and B1 DNA signals in the nucleus. The FISH segregation index is lowest in M-phase cells, but increases significantly in the G1 phase and peaks in the S phase (Fig. 3b).

Fig. 3: Dynamic segregation of L1 and B1 compartments during the cell cycle and embryonic development.
figure 3

a, b DNA FISH analysis of synchronized mESCs at different cell cycle stages. Representative images and the scatterplot of the segregation index of L1 and B1 DNA are shown in a and b, respectively. Data are presented as means ± standard deviation (SD). n, number of nuclei analyzed. M phase and G1 phase data are compared to S phase using the two-tailed Student’s t-test. P values are shown at the top. c Boxplot analysis of the ratio of homotypic interactions versus heterotypic interactions for B1 and L1 based on single-cell Hi-C data from mESCs. The y-axis shows the segregation index (Hi-C) of L1 and B1, which represents the ratio of average interaction frequency between compartments containing similar repeats (B1.B1 and L1.L1) to that between compartments containing different repeats (B1.L1) for all chromosomes (except X and Y). Larger values indicate a higher degree of homotypic interaction between B1- or L1-rich compartments. P values are calculated with the two-tailed Student’s t-test. d Heatmaps of normalized interaction frequencies at 500-kb resolution on chromosome 17 in mESCs at M (left) and G1 (middle) phase. A comparison of contact frequencies between M and G1 phase [log2(G1/M)] for the whole of chromosome 17 is shown on the right. e, f DNA FISH analysis of early embryos. Representative FISH images and the scatterplot of the segregation index of L1 and B1 DNA are shown in e and f, respectively. Data are presented as means ± SD (embryos were collected and processed in two independent experiments). n, number of nuclei analyzed. Each sample is compared to sample at the blastocyst stage and P values are calculated with the two-tailed Student’s t-test. Dotted lines show the trend-line. g Heatmaps of normalized interaction frequencies at 500-kb resolution on chromosome 17 in mouse embryos at the early 2-cell, late 2-cell, 8-cell, inner cell mass (ICM) stages and also in embryos treated with the transcription inhibitor α-amanitin for 20 and 45 h. Genomic densities of B1 and L1 repeats are shown at the bottom. h Boxplots showing the Hi-C segregation index of L1 and B1 in early embryos at various stages. The first six samples show a gradual increase of homotypic versus heterotypic interactions between L1-rich and B1-rich regions from zygotes (including PN3 and PN5 stages) to 8-cell embryos and pluripotent mESCs (in vitro equivalent of the inner cell mass cells of blastocysts). Each sample is compared to mESCs and P  values are calculated with the two-tailed Student’s t-test. The last two samples show the segregation index for embryos treated with α-amanitin for 20 or 45 h. P values are calculated with two-tailed Student’s t-test.

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To provide molecular evidence for the segregation of repeats during the cell cycle, we analyzed the published Hi-C data from cell-cycle synchronized mESCs and HeLa cells.78,79 In both cell types, G1-phase cells exhibit a classic plaid pattern of hierarchical interactions, with enriched and depleted interaction blocks outside of the diagonal region of the Hi-C interaction heatmap (Fig. 3c, d and Supplementary information, Fig. S6c, d). In contrast, M-phase cells exhibit stronger signals along the diagonal, which represent the linearly organized, longitudinally compressed array of consecutive chromatin loops. To quantify this difference, we defined a Hi-C-based segregation index by calculating the ratio of homotypic versus heterotypic interaction frequencies between L1 and B1/Alu subfamilies. Indeed, the Hi-C segregation index is significantly higher in G1-phase cells than in M-phase cells (Fig. 3c and Supplementary information, Fig. S6c). These results indicate that segregation of B1/Alu and L1 repeats is dispersed by mitosis, and is re-established when the higher-order chromatin structure forms during each cell cycle in both mouse and human cells. This finding agrees with previous reports that in metaphase, chromosome folding becomes homogeneous and large megabase-scale A and B compartments are lost, whilst in interphase, chromosomes return to a highly compartmentalized state.78,79

Dynamic establishment of L1 and B1 segregation in early embryogenesis

After fertilization, the chromatin undergoes extensive reprogramming from a markedly relaxed state in zygotes to fully organized structures in blastocysts.80,81,82 We performed a time-course DNA FISH analysis of L1 and B1 in early mouse embryos. During embryonic divisions, L1 and B1 signals are largely overlap** in zygotes, and become progressively more segregated in 2-cell, 4-cell, morula and blastocyst embryos (Fig. 3e, f). Consistently, analysis of the published Hi-C data of early embryos81 showed that early 2-cell embryos exhibit prevalent cis-chromosomal contacts along the diagonal of the Hi-C interaction map, whereas the plaid patterns of Hi-C interactions become readily detectable in late 2-cell embryos and are fully established in the inner cell mass (ICM) cells of blastocysts (Fig. 3g and Supplementary information, Fig. S7a). Plotting the FISH and Hi-C segregation indexes showed a gradual increase of L1 and B1 segregation along the course of blastocyst development, which reaches the highest level in blastocysts or in mESCs (Fig. 3f, h). We conclude that, in early embryos, compartmentalization of L1- and B1-rich regions appears to be established in a stepwise manner, coincident with de novo establishment of higher-order chromatin structures. Notably, the greatest change (steepest trend-line) of FISH segregation indexes occurs between the zygote and the late 2-cell stage (Fig. 3f), which implies that the initiation of B1 and L1 compartmentalization may coincide with the zygotic genome activation, during which massive transcription switches on.

It was reported that inhibition of Pol II by α-amanitin caused embryonic arrest at the late 2-cell stage,83 yet the higher-order chromatin structure could still be established.80,81 However, compared to the control groups, we found that α-amanitin-treated embryos exhibited significantly lower L1/B1 segregation indexes and less clear patterns of Hi-C plaids (Fig. 3g, h). At 20 h, α-amanitin-treated embryos showed a low median level of L1/B1 segregation indexes and a Hi-C pattern with extensive diagonal signals that are similar to those of early 2-cell embryos, while the control group had proceeded into the late 2-cell stage (Fig. 3g, h). At 45 h, the control group had proceeded into the 8-cell and morula stages, whereas α-amanitin-treated embryos (45 h) still showed low L1/B1 segregation and a Hi-C plaid pattern similar to that of late 2-cell embryos, despite the segregation index modestly increases compared with that of embryos at 20 h (Fig. 3g, h and Supplementary information, Fig. S7b). These results indicate a delayed and incomplete formation of the higher-order chromatin structure in the absence of zygotic Pol II transcription in mouse.

In accordance with delayed chromatin folding in embryos, treatments of mESCs with the drug 5,6-dichloro-1-β-d-ribofuranosylbenzimidizole (DRB) which inhibits Pol II transcription elongation, led to a partial loss of L1 perinucleolar localization and a gain of mixed nuclear L1 and B1 signals (Fig. 4a, b and Supplementary information, Fig. S7c). Inhibition of both Pol I and II by a high concentration of actinomycin D (ActD) had a more severe effect compared to DRB treatment (Fig. 4a, b). Thus, in both ESCs and early embryos, inhibition of Pol II transcription appears to partially, but not completely, block L1/B1 segregation. These results imply that L1/B1 compartmentalization is likely to be autonomously initiated and subsequently facilitated by transcription.

Fig. 4: Repeat RNA and transcription promote the spatial segregation of L1 and B1 compartments.
figure 4

a, b DNA FISH analysis of L1 (green) and B1 (red) repeats in mESCs treated with transcription inhibitors for 3 h. Representative images and the scatterplot of the segregation index of L1 and B1 DNA are shown in a and b, respectively. DMSO, mock control; DRB (100 μM), a drug inhibits the release and elongation of Pol II; ActD (1 μg/mL), a drug inhibits both Pol I and II. Both drug treatments disrupt the perinucleolar staining of L1 DNA and induce mixing of the L1 and B1 DNA signals. Treatment with ActD elicits a stronger mixing effect than DRB. c Developmental analysis of embryos microinjected with scramble (SCR) or L1 AMO (panel (i)) or with SCR or two different B1 ASOs (ASO-1 and ASO-2) (panel (ii)) at the zygote stage. n, number of embryos analyzed. d Embryos depleted of L1 RNA by AMO or B1 RNA by ASO show poorer L1/B1 segregation, as indicated by significantly lower L1/B1 segregation indexes compared to the scramble AMO/ASO controls and noninjected late 2-cell embryos. Each dot represents an embryo analyzed. Data were collected from three independent experiments. P values are calculated with the two-tailed Student’s t-test. e, f DNA FISH analysis of L1 (green) and B1 (red) repeats in mESCs transfected with scramble AMO for 36 h, or L1 AMO for 12 and 36 h, or treated with the drug AZT for 12 h. Representative images and the scatterplot of the segregation index of L1 and B1 DNA are shown in e and f, respectively. All scale bars, 5 μm. n, number of nuclei analyzed. Data are presented as means ± SD (> 3 independent experiments, except two biological replicates for AZT treatment), and P values are calculated with two-tailed Student’s t-test.

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Repeat transcripts promote L1 and B1 segregation in embryonic cells

In an effort to link repeat function with chromatin structure, we sought to explore the role of repeat RNA that is transcribed from L1 and B1 sequences. Both L1 and B1 repeats are activated and highly expressed in two-cell embryos (Supplementary information, Fig. S7d).71,84,85,86 We have reported previously that depletion of L1 RNA by an antisense morpholino (AMO) in mouse embryos led to arrest at the 2-cell stage; and in mESCs, its depletion led to reduced proliferation and global de-repression of hundreds of L1-associated genes; however, it did not alter the expression of OCT4 and NANOG, two known master regulators of the pluripotency program, nor induced ESC differentiation.61,87 Using the same L1 AMO sequence, we depleted L1 RNA by 17.4% on average shown by RNA FISH (n = 16 embryos; Supplementary information, Fig. S8a). This modest depletion is consistent with the general consensus that AMO acts through steric blockage of its target RNA rather than inducing RNA degradation. In concordance with the previous report by Percharde et al.,84 more than 91.3% of embryos (42 out of 46) were arrested at the 2-cell stage in contrast to only 15.4% of embryos treated with scramble AMO that were arrested (Fig. 4c), indicating effective inhibition of L1 RNA.

We also sought to perturb B1 expression by microinjecting B1 antisense oligonucleotides (ASO) into mouse zygotes. Two B1 ASOs significantly downregulated B1 RNA levels by 36% shown by RNA FISH (n = 17 embryos; Supplementary information, Fig. S8b). Strikingly, embryos depleted of B1 RNA were able to pass the first embryonic division, but failed to divide further and became arrested at the 2-cell stage (n = 79 embryos; Fig. 4c), indicating an essential requirement for B1 RNA in embryonic development. We collected these embryos for DNA FISH analysis when the control group injected with scramble AMO or ASO had grown to the late 2-cell stage. Compared to the control embryos, both L1- and B1-depleted embryos exhibited significantly lower L1/B1 FISH segregation indexes (Fig. 4d and Supplementary information, Fig. S8c, d), indicating delayed segregation of L1 and B1 compartments.

In order to dissect the effects independent of embryonic progression, we then tried to deplete B1 and L1 transcripts in mESCs. B1/Alu repeats have been broadly implicated in diverse processes, including transcription, RNA processing, and nuclear export.88,89,90,91 Treatment of mESCs with B1 ASO led to severe cell death within hours of transfection (data not shown), which precluded direct assessment of B1 RNA in chromatin organization. It has been suggested that nuclear organization is critically dependent on interactions within heterochromatin,34,92 where L1, the most abundant one of all repeat subclasses, is predominantly enriched. In subsequent analysis, we in-depth characterized the effects of depleting L1 RNA on chromatin organization.

Treatments of mESCs with L1 AMO led to a depletion of L1 RNA by 28% (n = 30 cells) (Supplementary information, Fig. S8e, f). At both 12 and 36 h post transfection of L1 AMO, nucleoplasmic signals of L1 DNA were obviously increased and perinucleolar L1 signals became fuzzy or absent (Fig. 4e). B1 signals became more uniformly dispersed in the nucleoplasm, in contrast to punctate staining of B1 in the control mESCs. Emergence of the overlap** L1 and B1 FISH signals is indicative of decreased homotypic clustering and segregation of L1 and B1 DNA. In comparison, treatment with the drug azidothymidine (AZT), which blocks L1 retrotransposition activity,S13a). For comparison, we also generated two synthetic DNA and RNA sequences in 1-kb length, comprising 8 tandem copies of either B1 element or scrambled B1 (designated as 8× B1 or 8× SCR, respectively). L1 mix as well as two synthetic fragments in DNA or RNA efficiently pulled down recombinant HP1α (Supplementary information, Fig. S13b). In addition, L1 RNA–HP1α interactions were robustly detected in highly stringent conditions with up to 1 M of salt and urea (Fig. 7e). These results indicate that HP1α exhibits strong binding activities towards RNA and DNA in vitro.

Recent studies have reported that HP1α forms phase-separated droplets in the presence of DNA or nucleosomes in vitro, and heterochromatin formation may entail a phase-separation mechanism.37,38,100,101,102 Indeed, the L1 DNA mix as well as two synthetic DNA controls (8× B1 and 8× SCR) promote the phase separation of HP1α, which fails to phase separate on its own (Supplementary information, Fig. S13c, d). Consistent with its strong RNA-binding activity, HP1α also phase separates with the L1 RNA mix in a concentration-dependent manner to form spherical droplets with liquid-like properties, such as fusion of droplets and rapid recovery after photo-bleaching (Fig. 7f and Supplementary information, Fig. S13d–f). Intriguingly, L1 RNA fragments from F3 to F6 with low GC contents (< 40%) covering the inter-ORF and central conserved ORF2 sequence (3.2-kb) tend to have higher activities in promoting HP1α droplet formation compared to F1 and F8 fragments with GC contents of 45%–56% (Supplementary information, Fig. S13g, h). In addition, careful examination of HP1α ChIP-seq in mESCs showed that HP1α preferentially binds to the central region of L1 repeat DNA (Supplementary information, Fig. S13i), which hints some degree of specificity in L1–HP1α interplay. However, there was no obvious difference detected between L1 mix and 8× B1 (60% in GC) RNA/DNA in HP1α phase separation (Supplementary information, Fig. S13j). Although we cannot conclude a sequence specificity of HP1α, clearly, HP1α shows strong RNA- and DNA-binding activities and phase-separates in the presence of RNA and DNA in vitro. Given the abundance and co-residence of L1 repeat DNA and RNA with HP1α within the nucleus, it is tempting to speculate that their co-localization may provide a means of specificity for L1 in promoting HP1α phase separation during heterochromatin formation.

Discussion

Although tremendous efforts have been dedicated to studies of structural chromatin proteins and cataloging chromatin maps, the role of DNA sequences in 3D genome organization has been largely ignored. Interestingly, the overall higher-order chromatin structure has been reported to be stable across different cell types and conserved in related species, despite occasional compartment switches in a small portion of the genome in a given cell (Supplementary information, Text S1). This remarkable conservation of chromatin compartments suggests a fundamental principle which all cells stick to while co** with shifting signals in different cell fates. Compared to transcription and epigenetic modifications, the primary DNA sequence has an unparalleled advantage to directly control and govern the stability of 3D genome folding due to its static nature during development. Then, the question comes what DNA sequences serve such a task as the blueprint of 3D genome folding. By employing in silico polymer simulations to interpret microscopy and Hi-C data, Mirny and colleagues have suggested that compartmental segregation may occur through a microphase-separation mechanism of block copolymers.29,34 Together with the Solovei group, they further proposed that interactions between heterochromatic regions, rather than euchromatic contacts and lamina-heterochromatin interactions, are crucial for compartmentalization of the genome in both inverted and conventional nuclei.29,34 However, what was unknown in their model is the molecular determinants, particularly the genetic information of block copolymers that are responsible for chromatin compartmentalization.

In this study, we reveal a remarkable correlation between repeat distribution and compartmental organization of the higher-order chromatin structure. First, using complementary genomics and imaging approaches, we demonstrate that the self-clustering of L1 and B1/Alu repeats forms grossly exclusive nuclear domains that are highly correlated with and predict the known A/B compartments, and that nuclear segregation of L1-rich and B1/Alu-rich sequences is conserved across mouse and human cells, and can be dynamically established during cell division and early embryogenesis. Second, we show that depletion of L1 RNA by AMOs drastically alters repeat segregation and compartmentalization on a global scale and at individual loci by Hi-C, DNA FISH and Oligopaint FISH. Collectively, the overall positive correlation and the essentiality of L1 RNA in compartmental organization suggest a functional role for L1 repeats in driving genome folding. These results disfavor the notion that L1 or B1/Alu repeats are merely markers of large chromosomal segments with a different activity, although we cannot firmly exclude this possibility. Our model is also consistent with the growing evidence showing active roles of retrotransposons in re-wiring the genome and regulatory programs.53,54,55,56,57,58,59,60,61 As often a challenge for genome organization studies, we note that the current support for going beyond correlative evidence to really show a causative role of L1 is still limited.

L1 RNA tends to co-localize with L1 DNA sequences in regions enriched for the binding of HP1α and H3K9me3. Intriguingly, L1 RNAs can also be detected outside the nuclear and nucleolar periphery (Supplementary information, Fig. S12b). L1 RNA has a short half-life of 40 min.61 Torres-Padilla and colleagues reported previously that exogenous L1 RNA fails to rescue the chromatin defects upon abnormal silencing of L1 in mouse zygotes.71 Although these observations disfavor a trans-acting mechanism, it is possible that L1 transcripts might be mobilized to distal L1 DNA sequences. As 20%–40% of L1 repeats are located in annotated euchromatic compartments (Supplementary information, Fig. S1e), these L1 transcripts may be more readily visualized by microscopy. For the majority of L1 enriched in transcriptionally silenced heterochromatic environments, their expression might be temporally regulated in a more transient way, thus creating difficulty for direct visualization and detection. Studies of X chromosome inactivation revealed different roles for silenced and actively transcribed L1s in regulating heterochromatin formation induced by ** by RNA in gene regulatory elements. Science 350, 978-981 (2015)." href="/article/10.1038/s41422-020-00466-6#ref-CR105" id="ref-link-section-d206178417e2522">105), or acting as a scaffold to anchor L1-rich chromosomal segments to the nucleolar and nuclear peripheries. Cases for RNA in organizing subnuclear domains have been reported. For example, ** of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009)." href="/article/10.1038/s41422-020-00466-6#ref-CR6" id="ref-link-section-d206178417e2761">6 and compartmentalization strength23,34,94 were identified as described previously.

Segregation index

The FISH-based segregation index is defined as the negative value of Pearson’s correlation coefficient of L1 and B1 DNA signals in the nucleus. The Hi-C-based segregation index is defined as the ratio of homotypic versus heterotypic interaction frequencies between L1 and B1/Alu subfamilies.

In vitro pull-down and phase separation assays

A series of eight 1-kb fragments, designated as F1 to F8, were produced to cover the full-length 6544-kb L1 sequence by PCR (Supplementary information, Fig. 13a). Two artificial fragments comprising eight tandem copies of either B1 element (8× B1, in 1-kb length) or scrambled B1 sequence (8× SCR, in 1-kb length) were used for comparison. Biotin-labeled RNA was obtained by in vitro transcription for the pull-down experiment. We purified the recombinant human HP1α protein as previously described.38 In phase separation assays, recombinant HP1α with DNA or RNA fragments for L1, 8× B1, or 8× SCR were incubated at 4 °C overnight.

Statistical analysis

Statistical analyses were carried out using Excel or R (version 3.4.3).

Please also see Supplementary information, Data S1 for details.