Constitutive heterochromatin is an important component of eukaryotic genomes that has essential roles in nuclear architecture, DNA repair and genome stability1, and silencing of transposon and gene expression2. Heterochromatin is highly enriched for repetitive sequences, and is defined epigenetically by methylation of histone H3 at lysine 9 and recruitment of its binding partner heterochromatin protein 1 (HP1). A prevalent view of heterochromatic silencing is that these and associated factors lead to chromatin compaction, resulting in steric exclusion of regulatory proteins such as RNA polymerase from the underlying DNA3. However, compaction alone does not account for the formation of distinct, multi-chromosomal, membrane-less heterochromatin domains within the nucleus, fast diffusion of proteins inside the domain, and other dynamic features of heterochromatin. Here we present data that support an alternative hypothesis: that the formation of heterochromatin domains is mediated by phase separation, a phenomenon that gives rise to diverse non-membrane-bound nuclear, cytoplasmic and extracellular compartments4. We show that Drosophila HP1a protein undergoes liquid–liquid demixing in vitro, and nucleates into foci that display liquid properties during the first stages of heterochromatin domain formation in early Drosophila embryos. Furthermore, in both Drosophila and mammalian cells, heterochromatin domains exhibit dynamics that are characteristic of liquid phase-separation, including sensitivity to the disruption of weak hydrophobic interactions, and reduced diffusion, increased coordinated movement and inert probe exclusion at the domain boundary. We conclude that heterochromatic domains form via phase separation, and mature into a structure that includes liquid and stable compartments. We propose that emergent biophysical properties associated with phase-separated systems are critical to understanding the unusual behaviours of heterochromatin, and how chromatin domains in general regulate essential nuclear functions.
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Funding was provided by National Institute of Health grants R01 GM117420 (G.H.K.), R01-GM074233 (D.V.F.), and U54-DK107980 and U01-EB021236 (X.D.), and the California Institute for Regenerative Medicine (CIRM, LA1-08013, X.D.). We thank A. Dernburg for use of the spinning disk confocal, M. Scott for maintenance of the rastering confocal, S. Colmenares for cell lines, C. Robertson for introduction to RICS and N&B, A. Tangara for help in assembling and maintaining the LLSM, and A. Dernburg, S. Safran, M. Elbaum and the Karpen laboratory for helpful comments on the manuscript.
The authors declare no competing financial interests.
Reviewer Information Nature thanks E. Selker, K. Rippe and the other anonymous reviewer(s) for their contribution to the peer review of this work.
Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended data figures and tables
a, Analysis of HP1a 206 amino acid protein sequence. Top, known domains chromo, hinge, and shadow. Black line, predictor of natural disordered regions (PONDR) score for intrinsic disorder, >0.5 is considered disordered. Red line, hydropathy score (positive is hydrophobic). Yellow bars indicate low-complexity sequences. b, 1 mg ml−1 HP1a in 50 mM NaCl was incubated at 37 °C for 5 min, then returned to room temperature (22 °C) and imaged with differential interference contrast microscopy (DIC) every 5 s for 8 min. Quantification of average number and area of HP1a droplets formed in a 50 × 50 μm window, n = 3. c, Probability distribution of droplet aspect ratio. d, Probability distribution of droplet area on a log–log plot follows a power law exponential with τ = −1.5, characteristic of aggregating systems. e, f, Sediment gradient analysis shows large oligomers of HP1a in 0.05 M NaCl (arrows, e) but not 0.5 M NaCl (f). g, Two-colour images showing HP1a and H2Av for one nucleus over Drosophila embryonic cycle 13. h, i, Quantification of average HP1a domain size per nucleus (sum of all foci in each nucleus) (h) and total intensity of fluorescently-tagged HP1a and H2A (i), over embryonic cycles 10–14. Error bars are s.d. n = 12 embryos of >75 nuclei each. j, Two-colour images of HP1a and H2A showing differentially shaped heterochromatin domains in Drosophila embryos, adult gut and cultured Kc cells, and mouse fibroblast NIH3T3 cells.
a, b, FRAP images (a) and average fluorescence intensity over time of bleached area (b) of Drosophila embryonic nuclei in cycles 10–14, and stage 8. In cycle 14, heterochromatin forms in the apical region of the nucleus. n = 20 nuclei in each condition, error bars are s.d. c, NIH3T3 nuclear size after addition of media containing 10% 1,6-hexanediol. n = 63 nuclei, error bars are s.d. d, Images and quantification of FRAP on wild-type (WT) HP1a and point mutants incapable of dimerizing (I191E) or interacting with binding partners (W200A) in S2 cells. n = 60 nuclei each condition, error bars are s.d.
a, Schematic illustration of dynamic properties near a phase boundary. Orange particles have self-association properties (green dotted lines) and concentrate into a phase (orange background, right side). An orange particle in the orange phase can move in any direction and encounter only orange particles (eight arrows) until it contacts a blue particle, which prevents self-association between two orange particles and limits the potential diffusion dimensions of the orange particle (red squiggle, loss of arrows). This results in two properties of particles near the phase boundary: net slower diffusion and higher likelihood that two orange particles will move in the same direction. b, Schematic illustrating number and brightness technique22. If particles are moving independent of one another (left, blue), variations in fluorescence intensity measured in the pixel volume over time will be small. If particles are moving together in and out of the pixel volume (right, red), intensity variation will be larger. This can result from bound molecules (that is, complex) or unbound molecules moving in the same direction (coordinated movement). c, A cultured Drosophila S2 cell expressing GFP-tagged fibrillarin to mark the nucleolus (left). Inset shows entire nucleus (dotted line) and region of interest (white box). Visual representation of increased variance measured by number and brightness (middle), colour scale represented in quantification graph (right). Quantification of variance (right) across the border from nucleoplasm to nucleolus (example line drawn in middle). Dotted line represents approximate nuclear boundary. d–g, Image, variance map, and quantified variance of HP4 in S2 cells (d), HP5 in S2 cells (e), HP1γ in mammalian NIH3T3 cells (f), and HP1c in S2 cells (g). h, Diffusion rate (D) for fibrillarin across the nucleoplasm–nucleolus boundary (left), and HP4 (middle) and HP5 (right) across the euchromatin–heterochromatin boundary. For c–h, n = 25 nuclei, error bars are s.d. i, Representative image, variance map, and quantified variance across boundary for HP4 in control cells (bw RNAi, top), HP1a-depleted cells (HP1a RNAi, middle), and HP1a-depleted and rescued cells (HP1a RNAi + mCherry–HP1a, bottom). n = 25 nuclei per condition, error bars are s.d. j, Predictions of inert probe variance and diffusion near ‘H2A edges’, which have a >2 fold increase in H2A density (purple bar) with <1.25 fold change in HP1a density (green bar), and ‘HP1a edges’, which have >3 fold increase in HP1a density (green bar) with <1.25 fold change in H2A density (purple bar). The chromatin compaction model (left) predicts that inert probe variance and diffusion rate would be influenced by increasing H2A density, but unaffected by HP1a density. The phase-separation model (right) predicts that inert probe variance and diffusion rate would be influenced by HP1a density, but unaffected by H2A density. k, Summary of RICS and N&B data. Heterochromatin proteins can move freely in the heterochromatin domain (i) but are hindered near the hetero-euchromatic border (ii). Similarly, euchromatic proteins move freely in euchromatin (iii) but are hindered near the border (iv), mostly preventing their entry. Euchromatic proteins that do enter heterochromatin move more slowly owing to energetically costly interactions with surrounding phase particles (v) or crowded environments (vi). l, Models of how liquid properties could influence heterochromatin domain formation and functions. We speculate that nucleation of heterochromatic (HP1) foci could occur independently of chromatin and H3K9me2/3, then associate with the chromatin fibre (top left), or nucleation could require H3K9me2/3 (top right). Heterochromatin could spread along the chromatin fibre (in cis) due to HP1 liquid ‘wetting’, followed by H3K9 HMTase recruitment and H3K9 methylation (‘HP1 first’), or as previously proposed, HP1 recruitment of the HMTase could result in H3K9 methylation of adjacent nucleosomes, followed by HP1 binding (H3K9me2/3 first). Finally, non-contiguous segments of chromatin (on the same or different chromosomes) can coalesce into one 3D domain, owing to liquid-like fusion events between H3K9me2/3–HP1-enriched regions.
HP1a-GFP embryo imaged on a lattice light sheet microscope over nuclear cycles 13-14. Note formation, growth, fusion and maturation of HP1a foci. A higher quality version of this video can be viewed here https://youtu.be/o3NV22f_8Wk
An xy view of one nucleus over cycle 13, imaged on a lattice light sheet microscope. A higher quality version of this video can be viewed here https://youtu.be/tqHV7hQrVME
An xz view of the same nucleus in Video S2 over cycle 13, imaged on a lattice light sheet microscope. A higher quality version of this video can be viewed here https://youtu.be/SUcAOTKXXVs
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Strom, A., Emelyanov, A., Mir, M. et al. Phase separation drives heterochromatin domain formation. Nature 547, 241–245 (2017). https://doi.org/10.1038/nature22989
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