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.

Many eukaryotic transcription factors (TFs) contain intrinsically disordered low-complexity sequence domains (LCDs), but how these LCDs drive transactivation remains unclear. We used live-cell single-molecule imaging to reveal that TF LCDs form local high-concentration interaction hubs at synthetic and endogenous genomic loci. TF LCD hubs stabilize DNA binding, recruit RNA polymerase II (RNA Pol II), and activate transcription. LCD-LCD interactions within hubs are highly dynamic, display selectivity with binding partners, and are differentially sensitive to disruption by hexanediols. Under physiological conditions, rapid and reversible LCD-LCD interactions occur between TFs and the RNA Pol II machinery without detectable phase separation. Our findings reveal fundamental mechanisms underpinning transcriptional control and suggest a framework for developing single-molecule imaging screens for drugs targeting gene regulatory interactions implicated in disease.

We imaged transcription in living cells using a locus-specific reporter system, which allowed precise, single-cell kinetic measurements of promoter binding, initiation and elongation. Photobleaching of fluorescent RNA polymerase II revealed several kinetically distinct populations of the enzyme interacting with a specific gene. Photobleaching and photoactivation of fluorescent MS2 proteins used to label nascent messenger RNAs provided sensitive elongation measurements. A mechanistic kinetic model that fits our data was validated using specific inhibitors. Polymerases elongated at 4.3 kilobases min −1 , much faster than previously documented, and entered a paused state for unexpectedly long times. Transcription onset was inefficient, with only 1% of polymerase-gene interactions leading to completion of an mRNA. Our systems approach, quantifying both polymerase and mRNA kinetics on a defined DNA template in vivo with high temporal resolution, opens new avenues for studying regulation of transcriptional processes in vivo.Transcription by RNA polymerase II (Pol II) is at the core of gene expression and hence is the basis of all cellular activities. Little information exists about the kinetics of this process in live cells 1 , as understanding of gene expression regulation comes from studies using purified proteins. For instance, the subunits of the elongating Pol II are well known 2 and the crystal structure of this enzyme explains much of its behavior in vitro 3,4 . mRNA transcription can be deconstructed into a succession of steps: promoter assembly, clearanceCorrespondence should be addressed to R.H.S. (rhsinger@aecom.yu.edu). AUTHOR CONTRIBUTIONS All data were initially acquired by X.D. and Y.S.-T. Subsequent data were obtained by V.d.T. (Fig. 4a,b and Fig. 5a) and Y.B. (Fig. 9b). S.M.S. was responsible for the microscopy, built the wide-field microscope for live-cell imaging and wrote analysis software. X.D. performed the kinetic modeling. R.D.P. provided consultation on model formulation and testing, and training in the use of the ProcessDB software. R.H.S. supervised the project. COMPETING INTERESTS STATEMENTThe authors declare competing financial interests: details accompany the full-text HTML version of the paper at http:// www.nature.com/nsmb/. HHS Public AccessAuthor manuscript Nat Struct Mol Biol. Author manuscript; available in PMC 2016 July 12. Author Manuscript Author ManuscriptAuthor ManuscriptAuthor Manuscript and escape 5 , followed by elongation and termination. The process of transcriptional initiation involves several structural changes in the polymerase as the nascent transcript elongates 6 . Early in initiation, the polymerase can produce abortive transcripts 7,8 . These abortive cycles have been observed with a single prokaryote polymerase (RNAP) releasing several transcripts without escaping the promoter 9,10 . The elongation step can be regulated by pausing for various times, as demonstrated using prokaryotic polymerases in vitro 11,12 .For eukaryotic cells, attempts have been mad...

Folding of mammalian genomes into spatial domains is critical for gene regulation. The insulator protein CTCF and cohesin control domain location by folding domains into loop structures, which are widely thought to be stable. Combining genomic and biochemical approaches we show that CTCF and cohesin co-occupy the same sites and physically interact as a biochemically stable complex. However, using single-molecule imaging we find that CTCF binds chromatin much more dynamically than cohesin (~1–2 min vs. ~22 min residence time). Moreover, after unbinding, CTCF quickly rebinds another cognate site unlike cohesin for which the search process is long (~1 min vs. ~33 min). Thus, CTCF and cohesin form a rapidly exchanging 'dynamic complex' rather than a typical stable complex. Since CTCF and cohesin are required for loop domain formation, our results suggest that chromatin loops are dynamic and frequently break and reform throughout the cell cycle.DOI: http://dx.doi.org/10.7554/eLife.25776.001

Materials and MethodsMicro-C protocol for mammals was modified from the original protocol for yeast in (1, 2). The protocol was optimized for the input cell number from 1k to 5M and first applied to the mammalian system in (3). We first briefly summarize the critical steps and concepts in the Micro-C method, and then provide detailed step-by-step instructions. Micro-C experiment 1. Cell culture and crosslinkingHere, we performed a dual crosslinking protocol to fix protein-DNA and protein-protein interactions. In addition to formaldehyde, we used the non-cleavable and membrane-permeable protein-protein crosslinker DSG (disuccinimidyl glutarate, 7.7Å) or EGS (ethylene glycol bis(succinimidyl succinate), 16.1Å) to crosslink the primary amines between proximal proteins. The dual-crosslinking method significantly increases the signal-to-noise ratio of Micro-C data in yeast (2).In brief, 1k -5M cells were resuspended by trypsin and fixed by freshly made 1% formaldehyde at room temperature for 10 minutes. The crosslinking reaction was quenched by adding Tris buffer (pH = 7.5) to final 0.75 M at room temperature. Fixed cells were washed twice with 1X PBS and protein-protein interactions fixed by 3 mM DSG for 45 minutes at room temperature. The DSG solution was freshly made at a 300 mM concentration in DMSO and diluted to 3 mM in 1X PBS before use. The crosslinking reaction was quenched by 0.75 M Tris buffer and washed twice with 1X PBS. Crosslinked cells were snap-frozen in liquid nitrogen and stored at -80°C (pellets are stable for up to a year). Note that freshly made crosslinking solution is critical to producing high-reproducibility Micro-C data, and Tris buffer is a faster and stronger quenching agent than glycine.