Chromatin is reprogrammed after fertilization to produce a totipotent zygote with the potential to generate a new organism1. The maternal genome inherited from the oocyte and the paternal genome provided by sperm coexist as separate haploid nuclei in the zygote. How these two epigenetically distinct genomes are spatially organized is poorly understood. Existing chromosome conformation capture-based methods2,3,4,5 are not applicable to oocytes and zygotes owing to a paucity of material. To study three-dimensional chromatin organization in rare cell types, we developed a single-nucleus Hi-C (high-resolution chromosome conformation capture) protocol that provides greater than tenfold more contacts per cell than the previous method2. Here we show that chromatin architecture is uniquely reorganized during the oocyte-to-zygote transition in mice and is distinct in paternal and maternal nuclei within single-cell zygotes. Features of genomic organization including compartments, topologically associating domains (TADs) and loops are present in individual oocytes when averaged over the genome, but the presence of each feature at a locus varies between cells. At the sub-megabase level, we observed stochastic clusters of contacts that can occur across TAD boundaries but average into TADs. Notably, we found that TADs and loops, but not compartments, are present in zygotic maternal chromatin, suggesting that these are generated by different mechanisms. Our results demonstrate that the global chromatin organization of zygote nuclei is fundamentally different from that of other interphase cells. An understanding of this zygotic chromatin ‘ground state’ could potentially provide insights into reprogramming cells to a state of totipotency.
To investigate 3D genome organization in nuclei of single cells, we developed a genome-wide high-resolution in situ Hi-C approach. Conventional Hi-C methods include biotin incorporation and enrichment for ligated fragments6, which might limit fragment retrieval. We simplified the protocol by omitting these steps, similarly to genome conformation capture7 (Fig. 1a, Extended Data Fig. 1, see Supplementary Methods). To verify the protocol, we compared data from population and single-cell data from K562 (human chronic myelogenous leukaemia) cells and obtained a contact probability Pc(s), dependent on genomic distance, s, that matched conventional in situ Hi-C on bulk K562 cells8 (Fig. 1b). When applied to oocytes (Fig. 1c), our method was efficient at capturing chromosomal interactions: single-nucleus Hi-C (snHi-C) revealed up to 1.9 × 106 contacts per cell after filtering, yielding 1–2 orders of magnitude more contacts than published single-cell Hi-C data2 and exceeding contact frequencies in a recent single-cell Hi-C preprint and report9,10. Half of the cells had >3.39 × 105 contacts per cell and 7.1% had >1 × 106 contacts per cell (Supplementary Table 1). These high-density snHi-C data enabled us to examine chromatin features directly in single-cell maps.
To investigate higher-level chromatin organization in oocytes, we examined how contact probability Pc(s)6,11 depends on genomic distance in individual cells and pooled data. In oocytes, the shapes of Pc(s) curves were consistent between individual cells (Fig. 1d) but markedly different from the characteristic shape in other mammalian interphase cells (Fig. 1e). For genomic separation s > 1 Mb, we observed steeper (~s−1.5) decay in oocyte Pc(s), closer to the random walks of yeast chromosomes12,13,14. Our simulations (see below) showed that steeper Pc(s) can be attributed to the larger volume of oocyte nuclei (~25 μm versus ≤10 μm diameter in somatic cells, Extended Data Fig. 2a).
Another major feature of mammalian chromosomes is segregation into active and inactive (A-B) compartments6. Although assignment of compartments from snHi-C data was impossible owing to its sparsity, an enrichment of interactions between the same compartment type and depletion between different types became evident in individual cells when compartments were assigned using GC content (Fig. 2a) or population Hi-C from other cell types (Extended Data Figs 3, 9). We also examined whether loops8 and TADs15,16, prominent functional features of chromatin organization17,18,19, are present in individual cells. Averaging over all genomic positions of loops and TADs identified in population Hi-C8 (for CH12-LX cells, see Supplementary Methods) revealed that both are present in individual oocytes as average enrichments (Fig. 2a, Extended Data Fig. 3b) but vary between cells (Extended Data Fig. 3c), reflecting both inter-cell variability and variations in experimental conditions. We conclude that a single nucleus shows enrichments of interactions between regions of the same compartment type, within TADs and at loops.
Using snHi-C data, we asked whether TADs constitute physically isolated domains in individual cells or reflect a mere tendency of chromosomes in a cell population to interact more within and less outside of a domain. We envisioned three scenarios (Fig. 2b): (i) all population-identified TADs are present in every individual cell; (ii) only population TAD boundaries are present, but individual TADs can be missing or fused in single cells; (iii) contacts may be clustered in individual cells, but clusters do not always match population TADs, revealing them only as an average feature. To distinguish between these scenarios, we examined a region in the snHi-C maps with most contacts (Fig. 2c, d) and segmented chromosomes into domains of enriched contact frequency (contact clusters) using an exact segmentation algorithm that maximizes modularity (see Supplementary Methods; comparable results for modularity segmentation of population TADs were obtained using an algorithm from ref. 20, see Extended Data Fig. 4a). We found that single-cell contact clusters do not always match population TADs, as contact clusters are highly variable and frequently occur across TAD borders. Nevertheless, variable contact clusters averaged into TADs when pooled together (Fig. 2d). The high cell-to-cell variations in contact patterns cannot be solely explained by experimental DNA loss because we often observe a presence of border-violating clusters rather than absence, and second, such patterns are also observed in regions of high read coverage (Extended Data Fig. 4b, c).
We validated frequent violations of TAD boundaries using 3D DNA fluorescence in situ hybridization (FISH) for equidistant pairs of probes located within a TAD and across a TAD border. Scenarios (i) and (ii), with TADs present in single cells as isolated domains, are expected to yield intra-TAD distances that are mostly shorter than inter-TAD distances. However, imaging of embryonic stem (ES) cells showed that inter-TAD distances are shorter than intra-TAD in 42% of cases, although the average inter-TAD distance is larger than the average intra-TAD (Wilcoxon test, P = 0.007). This indicates that although TAD borders affect the average distance and lead to preferential intra-TAD contacts, they cannot prevent contacts between TADs in single cells (Fig. 2e). Surprisingly, even the long inter-TAD pair was closer than the intra-TAD pair in 18% of cells, despite having twice the linear genome separation, having 50% larger average distance, and fourfold lower contact probability. Together, both imaging data and snHi-C support scenario (iii), in which TADs reflect a tendency for contact enrichment arising from a diverse conformational ensemble, rather than being isolated blocks of DNA present in individual cells.
The recently proposed mechanism of TAD formation by loop extrusion and boundary insulation21,22 provides a rationale for frequent boundary violations and, generally, TAD stochasticity. In this model, insulation prevents extruded loops from crossing boundaries, but it does not directly prevent contacts between TADs. Polymer simulations show that contact clusters naturally emerge from the 3D spatial proximity of DNA in a confined volume22 and frequently cross TAD boundaries (Extended Data Figs 5, 6 and ref. 11). As contact clusters are also detected in K562 snHi-C (Extended Data Fig. 4d), stochastic cluster formation is probably a universal property of chromatin organization in single cells.
Next, we investigated the spatial reorganization of chromatin during the transition from transcriptionally active immature oocytes (non-surrounded nucleolus, NSN) into transcriptionally inactive mature (surrounded-nucleolus, SN) oocytes23,24 (Fig. 3a). We observed a significant decrease in loop, TAD, and compartment strengths (Fig. 3b–d, Extended Data Fig. 7; all Mann–Whitney P < 0.005) during maturation, which may be related to transcriptional silencing and visual detachment of chromatin from the nuclear envelope23,24 (Fig. 3a). Whereas combined Pc(s) scalings are similar, mature oocytes display more long-range (>400 kb) contacts (Mood’s equal median test P = 0.02) and significantly less cell-to-cell variation in Pc(s) curves (Levene’s test, P = 0.007) (Fig. 3e–g). These findings are consistent with progressive chromatin reorganization during oocyte maturation.
We next addressed the key question, namely whether and how chromatin is reorganized during the oocyte-to-zygote transition, and whether it is different between the maternal and paternal genomes that have different biological histories and epigenetic modifications1. Oocyte chromosomes decondense after two meiotic divisions into the maternal nucleus. However, paternal nuclei are formed from compacted sperm by replacing protamines with histones25,26. To determine whether chromatin architecture is inherited or established de novo after fertilization, we extracted maternal and paternal nuclei from zygotes predominantly in the G1 phase and performed snHi-C (Figs. 1a, 4a; similar results were obtained without extracting nuclei, see Extended Data Fig. 8a, b and Supplementary Information). We detected up to 6 × 105 contacts in zygotic nuclei, which is twofold higher than in somatic cells and threefold lower than in the highest coverage oocytes. Results from averaging over TADs and loops identified previously8 showed that these features are present at similar strengths in maternal and paternal nuclei (Fig. 4b, Extended Data Figs 3, 9). Strikingly, although A–B compartmentalization is observed in paternal nuclei, it is notably absent from maternal nuclei (Fig. 4b, c, Extended Data Fig. 9a, b). To our knowledge, this is the first example of mammalian interphase nuclei presenting essentially no A–B compartmentalization. These results further suggest that the mechanisms forming compartments are distinct from those forming TADs and loops, in agreement with a recent preprint27.
To corroborate this novel finding, we simultaneously imaged 25 loci across chromosome 11 using 3D FISH and for each probe measured distances to the nearest probes of the same and of the different compartment type (Fig. 4d, Extended Data Fig. 10a, b). In agreement with Hi-C findings (Fig. 4c), we found that compartmentalization in ES cells is most pronounced (P < 10−16, one-sided Mann–Whitney U-test, Extended Data Fig. 10c); compartmentalization in paternal nuclei is weak but significant (P < 0.01); and compartmentalization in maternal nuclei is undetectable as compared to a randomized control with shuffled probe identities (P = 0.08, Fig. 4d, see Supplementary Methods). The lack of compartmentalization in the maternal genome may be due to a transcriptionally inactive extended G1 phase after fertilization1, suggesting that compartments are established de novo in the maternal genome1, whilst paternal genome compartmentalization is either inherited from sperm chromatin or established faster. The compartmentalization in the paternal nuclei aligns with detection of hyperacetylated histone H4, a hallmark of active chromatin, in early G1 phase and earlier transcriptional activation28.These results suggest a role for transcription in genome compartmentalization. We propose that the chromatin organization of zygotic nuclei denotes a ‘ground state’ produced by transcriptional silencing, chromosome condensation and an exchange in cohesin complex composition at fertilization29.
Finally, we used polymer modelling to understand global chromosome organization in the three analysed cell types and somatic cells. A prominent feature of Pc(s) curves is a steeper overall slope in oocytes and zygotes compared to somatic cells. Polymer modelling demonstrates that this steeper slope can be explained by the larger nuclear volumes of oocytes and zygotes (Extended Data Fig. 2). For maternal and paternal zygotic nuclei, experimental Pc(s) curves show similar shallow slopes for genomic distances s < 3 Mb (Fig. 4e), probably reflecting local compaction by loop extrusion that is also observed in simulations and that underlies formation of TADs and loops21,22. This scaling regime in zygotes is then followed by a plateau between 3–12 Mb for the maternal genome, whereas Pc(s) continues to decrease for the paternal genome. Simulations suggested that this difference may reflect previous states of differently compacted chromosomes in maternal and paternal zygotes. Simulations of decondensation subject to loop extrusion that start from a metaphase chromosome11 (Fig. 4g, Extended Data Fig. 6) result in Pc(s) that resemble those of maternal nuclei (Fig. 4h, Extended Data Fig. 2c). Analogous simulation starting from the compact fractal globule6, as a model of protamine-compacted state (Fig. 4g, Extended Data Figs 2b, 5), can reproduce paternal Pc(s) curve.
Taken together, these results suggest that the factors influencing Pc(s) are nuclear density, memory of the previous chromosome state, and cell cycle phase. Zygotic maternal nuclei and somatic cells are both predominantly in G1 phase and recently experienced chromosome decondensation from metaphase, which makes their global genome organization most similar. Paternal nuclei have a different biological history due to chromatin compaction by protamines in sperm and are thus different from somatic cells and maternal nuclei. Oocytes experienced the last mitosis weeks or months ago and are arrested in prophase I; they therefore differ the most from somatic cells.
In summary, our work provides insights into general principles of chromosome organization and specific biological aspects of oocyte and zygote genomes. We find that all known levels of chromosomal organization appear as mere tendencies that become visible when averaged over a population of cells or over loci; in single cells and at a single locus they may become overshadowed by the stochasticity of chromosome conformations. The unanticipated finding that zygotic maternal chromatin contains TADs and loops but not compartments suggests that it represents a transition towards building the embryonic chromatin organization of a totipotent cell. The difference in higher-order chromatin organization between maternal and paternal chromatin also raises the question of how paternal chromatin maintains or establishes compartments faster after fertilization. Our results, together with cohesin loader depletion experiments27, suggest that loops and compartments are formed by distinct mechanisms. Lastly, snHi-C could enable the study of chromatin organization during development and in rare cell types, such as stem cells and distinct cells within highly heterogeneous tumours. By combining snHi-C with other single-cell approaches, including single-cell transcriptome and methylome analyses, it will be possible to build a comprehensive picture of the interplay between genome folding and transcription in generating identities of individual cells.
Gene Expression Omnibus
We thank C. Theußl for help with pronuclear extraction procedure, S. Ladstätter for assistance in scoring oocyte stages and K. Klien for experimental support and mouse colony management. We are grateful to I. Adams, S. Boyle, I. Vassias-Jossic, G. Almouzni and W. Bickmore for advice and help with FISH experiments. Illumina sequencing was performed at the VBCF NGS Unit (http://www.vbcf.ac.at) except Hi-C libraries from MEL cells, which were sequenced in the Laboratory of Evolutionary Genomics of the Faculty of Bioengineering and Bioinformatics, Moscow State University, by M. Logacheva. K562 cells were a gift from Alexander Stark laboratory. We thank the staff of the Institute of Genetics and Molecular Medicine imaging facility and Vienna Biocenter BioOptics facility for assistance with imaging and analysis. We thank all members of the K.T.-K. laboratory for discussions, Life Science Editors for editorial assistance and R. Illingworth for critically reading the manuscript. J.G. is an associated student of the DK Chromosome Dynamics supported by the grant W1238-B20 from the Austrian Science Fund (FWF). H.B.B. was partly supported by the Natural Sciences and Engineering Research Council of Canada, PGS-D. This work was funded by the Austrian Academy of Sciences and by the European Research Council (ERC-StG-336460 ChromHeritance) to K.T.-K. as well as by a grant from the Russian Science Foundation (14-24-00022) to S.V.U. and S.V.R. The work in the Mirny laboratory is supported by R01 GM114190, U54 DK107980 from the National Institute of Health, and 1504942 from the National Science Foundation.
Extended data figures
This file contains Supplementary Methods, Supplementary References and Supplementary Table 1.
About this article
Nature Genetics (2019)