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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. A human cell contains about 2 meters of DNA tightly packed in a compartment called the nucleus. Within the space inside the nucleus, different parts of the DNA fold into distinct bundles known as domains. These domains are important for organising the genome and are crucial for regulating gene expression, by stimulating specific DNA segments to activate certain genes. Previous research has shown that DNA segments within the same domain frequently interact, whereas DNA segments in different domains rarely do. The domains are often folded into loops that are held together by a ring-shaped protein complex called cohesin, while another protein called CTCF positions cohesin and thereby sets the boundaries between the domains. Some mutations are known to disrupt these boundaries, which allows certain DNA segments to activate the wrong genes. This can lead to cancer or cause defects when embryos are developing. However, we do not currently understand how these domains are formed or maintained. In particular, it was unclear whether these loop domains are stable or dynamic structures. Hansen et al. addressed these questions in embryonic stem cells from mice and human cancer cells. It was found that cohesin and CTCF form a complex that binds to the DNA and likely holds the loops together. In further experiments, single molecules of cohesin and CTCF were tracked inside cells using super-resolution microscopy. The results showed that CTCF and cohesin bind to DNA with different dynamics: CTCF binds the DNA for about a minute, whereas cohesin binds the DNA for about 20–25 minutes. Once CTCF detaches from DNA, it quickly rebinds DNA at another site, but cohesin takes much longer. These observations suggest that rather than remaining static, chromatin domains are held together by a dynamic protein complex, with a molecular composition that exchanges over time. This results suggests that DNA loop domains, which were generally assumed to be very stable anchor points, are in fact highly dynamic structures that frequently fall apart and reform. The next challenge will be to understand how the dynamic nature of these loop domains contribute to gene regulation. This may, one day, enable us to manipulate the domains to correct faulty folding of DNA in cancer and other diseases.

Many components of eukaryotic transcription machinery—such as transcription factors and cofactors including BRD4, subunits of the Mediator complex, and RNA polymerase II—contain intrinsically disordered low-complexity domains. Now a conceptual framework connecting the nature and behavior of their interactions to their functions in transcription regulation is emerging (see the Perspective by Plys and Kingston). Chong et al. found that low-complexity domains of transcription factors form concentrated hubs via functionally relevant dynamic, multivalent, and sequence-specific protein-protein interaction. These hubs have the potential to phase-separate at higher concentrations. Indeed, Sabari et al. showed that at super-enhancers, BRD4 and Mediator form liquid-like condensates that compartmentalize and concentrate the transcription apparatus to maintain expression of key cell-identity genes. Cho et al. further revealed the differential sensitivity of Mediator and RNA polymerase II condensates to selective transcription inhibitors and how their dynamic interactions might initiate transcription elongation. Science , this issue p. eaar2555 , p. eaar3958 , p. 412 ; see also p. 329 Low-complexity domains of eukaryotic transcription factors form hubs via dynamic, multivalent, sequence-specific interactions. 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.

The enormous size of mammalian genomes means that for a DNA-binding protein the number of nonspecific, off-target sites vastly exceeds the number of specific, cognate sites. How mammalian DNA-binding proteins overcome this challenge to efficiently locate their target sites is not known. Here, through live-cell single-molecule tracking, we show that CCCTC-binding factor, CTCF, is repeatedly trapped in small zones that likely correspond to CTCF clusters, in a manner that is largely dependent on an internal RNA-binding region (RBR i ). We develop a new theoretical model called anisotropic diffusion through transient trapping in zones to explain CTCF dynamics. Functionally, transient RBR i -mediated trapping increases the efficiency of CTCF target search by ~2.5-fold. Overall, our results suggest a ‘guided’ mechanism where CTCF clusters concentrate diffusing CTCF proteins near cognate binding sites, thus increasing the local ON-rate. We suggest that local guiding may allow DNA-binding proteins to more efficiently locate their target sites. Single-particle tracking and mathematical modeling methods reveal the searching mechanism of CTCF for its cognate sites on DNA. An RNA-binding region in CTCF mediates its trapping in small zones and increases its target search efficiency.

The carboxy-terminal domain (CTD) of RNA polymerase (Pol) II is an intrinsically disordered low-complexity region that is critical for pre-mRNA transcription and processing. The CTD consists of hepta-amino acid repeats varying in number from 52 in humans to 26 in yeast. Here we report that human and yeast CTDs undergo cooperative liquid phase separation, with the shorter yeast CTD forming less-stable droplets. In human cells, truncation of the CTD to the length of the yeast CTD decreases Pol II clustering and chromatin association, whereas CTD extension has the opposite effect. CTD droplets can incorporate intact Pol II and are dissolved by CTD phosphorylation with the transcription initiation factor IIH kinase CDK7. Together with published data, our results suggest that Pol II forms clusters or hubs at active genes through interactions between CTDs and with activators and that CTD phosphorylation liberates Pol II enzymes from hubs for promoter escape and transcription elongation.

It remains unclear why acute depletion of CTCF (CCCTC-binding factor) and cohesin only marginally affects expression of most genes despite substantially perturbing three-dimensional (3D) genome folding at the level of domains and structural loops. To address this conundrum, we used high-resolution Micro-C and nascent transcript profiling in mouse embryonic stem cells. We find that enhancer–promoter (E–P) interactions are largely insensitive to acute (3-h) depletion of CTCF, cohesin or WAPL. YY1 has been proposed as a structural regulator of E–P loops, but acute YY1 depletion also had minimal effects on E–P loops, transcription and 3D genome folding. Strikingly, live-cell, single-molecule imaging revealed that cohesin depletion reduced transcription factor (TF) binding to chromatin. Thus, although CTCF, cohesin, WAPL or YY1 is not required for the short-term maintenance of most E–P interactions and gene expression, our results suggest that cohesin may facilitate TFs to search for and bind their targets more efficiently. Micro-C and nascent transcript profiling in mouse embryonic stem cells shows that enhancer–promoter interactions are robust to acute depletion of CTCF, cohesin, WAPL or YY1. Live-cell, single-molecule imaging shows that cohesin depletion reduces transcription factor binding to chromatin.

Single-particle tracking (SPT) directly measures the dynamics of proteins in living cells and is a powerful tool to dissect molecular mechanisms of cellular regulation. Interpretation of SPT with fast-diffusing proteins in mammalian cells, however, is complicated by technical limitations imposed by fast image acquisition. These limitations include short trajectory length due to photobleaching and shallow depth of field, high localization error due to the low photon budget imposed by short integration times, and cell-to-cell variability. To address these issues, we investigated methods inspired by Bayesian nonparametrics to infer distributions of state parameters from SPT data with short trajectories, variable localization precision, and absence of prior knowledge about the number of underlying states. We discuss the advantages and disadvantages of these approaches relative to other frameworks for SPT analysis.

Hyperphosphorylation of the C-terminal domain (CTD) of the RPB1 subunit of human RNA polymerase (Pol) II is essential for transcriptional elongation and mRNA processing 1 – 3 . The CTD contains 52 heptapeptide repeats of the consensus sequence YSPTSPS. The highly repetitive nature and abundant possible phosphorylation sites of the CTD exert special constraints on the kinases that catalyse its hyperphosphorylation. Positive transcription elongation factor b (P-TEFb)—which consists of CDK9 and cyclin T1—is known to hyperphosphorylate the CTD and negative elongation factors to stimulate Pol II elongation 1 , 4 , 5 . The sequence determinant on P-TEFb that facilitates this action is currently unknown. Here we identify a histidine-rich domain in cyclin T1 that promotes the hyperphosphorylation of the CTD and stimulation of transcription by CDK9. The histidine-rich domain markedly enhances the binding of P-TEFb to the CTD and functional engagement with target genes in cells. In addition to cyclin T1, at least one other kinase—DYRK1A 6 —also uses a histidine-rich domain to target and hyperphosphorylate the CTD. As a low-complexity domain, the histidine-rich domain also promotes the formation of phase-separated liquid droplets in vitro, and the localization of P-TEFb to nuclear speckles that display dynamic liquid properties and are sensitive to the disruption of weak hydrophobic interactions. The CTD—which in isolation does not phase separate, despite being a low-complexity domain—is trapped within the cyclin T1 droplets, and this process is enhanced upon pre-phosphorylation by CDK7 of transcription initiation factor TFIIH 1 – 3 . By using multivalent interactions to create a phase-separated functional compartment, the histidine-rich domain in kinases targets the CTD into this environment to ensure hyperphosphorylation and efficient elongation of Pol II. The histidine-rich domain of cyclin T1 promotes phase separation into liquid droplets, which facilitates the hyperphosphorylation of the C-terminal domain repeats of RNA polymerase II.

During mitosis, transcription is shut off, chromatin condenses, and most transcription factors (TFs) are reported to be excluded from chromosomes. How do daughter cells re-establish the original transcription program? Recent discoveries that a select set of TFs remain bound on mitotic chromosomes suggest a potential mechanism for maintaining transcriptional programs through the cell cycle termed mitotic bookmarking. Here we report instead that many TFs remain associated with chromosomes in mouse embryonic stem cells, and that the exclusion previously described is largely a fixation artifact. In particular, most TFs we tested are significantly enriched on mitotic chromosomes. Studies with Sox2 reveal that this mitotic interaction is more dynamic than in interphase and is facilitated by both DNA binding and nuclear import. Furthermore, this dynamic mode results from lack of transcriptional activation rather than decreased accessibility of underlying DNA sequences in mitosis. The nature of the cross-linking artifact prompts careful re-examination of the role of TFs in mitotic bookmarking. A kidney cell functions differently from a skin cell despite the fact that all the cells in one organism share the same DNA. This is because not all of the genes encoded within the DNA are active in the cells. Instead, cells can turn on just those genes that are specific to how that cell type works. One way that cells can regulate their genes is by using proteins called transcription factors that can bind to DNA to turn nearby genes on and off. When cells divide to form new cells, the DNA is condensed and gene activity is turned off. However, each dividing cell also has to ‘remember’ the program of genes that specifies its identity. After division, how do the cells know which genes to turn on and which ones to keep off? It was thought that the transcription factors attached to the DNA were all detached from it during cell division. Through studies in mouse embryonic stem cells, Teves et al. now show that this finding is largely an artifact of the methods used to study the process. In fact, many transcription factors still bind to and interact with DNA during cell division. This provides an efficient way for the newly formed cells to quickly reset to the pattern of gene activity appropriate for their cell type. Having found that many key transcription factors are still bound to DNA during cell division, the next challenge is to find out what role this binding plays in allowing cells to ‘remember’ their identity.

Single-particle tracking (SPT) has become an important method to bridge biochemistry and cell biology since it allows direct observation of protein binding and diffusion dynamics in live cells. However, accurately inferring information from SPT studies is challenging due to biases in both data analysis and experimental design. To address analysis bias, we introduce ‘Spot-On’, an intuitive web-interface. Spot-On implements a kinetic modeling framework that accounts for known biases, including molecules moving out-of-focus, and robustly infers diffusion constants and subpopulations from pooled single-molecule trajectories. To minimize inherent experimental biases, we implement and validate stroboscopic photo-activation SPT (spaSPT), which minimizes motion-blur bias and tracking errors. We validate Spot-On using experimentally realistic simulations and show that Spot-On outperforms other methods. We then apply Spot-On to spaSPT data from live mammalian cells spanning a wide range of nuclear dynamics and demonstrate that Spot-On consistently and robustly infers subpopulation fractions and diffusion constants. Proteins, the molecules that make up the cells’ internal machinery, are responsible for almost every process that keeps cells alive. Watching how proteins move and interact within a living cell can help scientists to better understand these biological mechanisms. Single-particle tracking is a recent technique that makes these observations possible by taking ‘live’ recordings of individual proteins in a cell. Typically, the goal of a single-particle tracking experiment is to assign proteins into groups, or subpopulations, based on the way they move in the cell. For example, one subpopulation may be bound to other cellular structures, a second moving freely at a high speed, and a third diffusing slowly. This informs on the biological roles of the proteins. The method involves an experimental stage and an analysis stage. During the experiment, proteins of interest are labeled with a small dye molecule that produces light when excited by a laser. The laser then illuminates the cell, stimulating all the labels in a thin layer. The position of each molecule is then determined with a microscope and a ‘snapshot’ taken. By repeating this process over multiple images, the movement of each molecule over time can be tracked. However, experimental problems can make the interpretation difficult. Motion blurring takes place when the proteins move so fast they appear as blurs in the images; tracking errors happen when so many proteins are present in the same space their trajectories overlap. Here, Hansen, Woringer et al. combine two pre-existing methods to improve the experimental set-up. Using lasers that flash like a strobe light reduces motion blurring by essentially taking snapshots of the proteins at short time intervals. Tracking errors are addressed by a technique whereby only one protein at a time produces light. Once the images are obtained and analyzed to yield trajectories, the trajectories themselves need to be analyzed to determine the number and properties of the protein subpopulations. Several factors can skew this analysis stage. For example, there is often a bias against fast-moving particles because the laser only lights up a thin layer of the cell. The proteins travelling slowly stay in focus long enough to be detected across many images; the fast ones quickly move out of the layer and are therefore counted less often. Hansen, Woringer et al. designed a free and user-friendly algorithm package called Spot-On to correct for this issue. Spot-On was thoroughly benchmarked against other solutions, demonstrating both its accuracy and robustness. Single-particle tracking can lead to misleading results if used incorrectly. It is essential to publically share solutions that help make this technique more rigorous, especially since a growing number of scientists have already started to use the method.

The regulation of transcription requires the coordination of numerous activities on DNA, yet how transcription factors mediate these activities remains poorly understood. Here, we use lattice light-sheet microscopy to integrate single-molecule and high-speed 4D imaging in developing Drosophila embryos to study the nuclear organization and interactions of the key transcription factors Zelda and Bicoid. In contrast to previous studies suggesting stable, cooperative binding, we show that both factors interact with DNA with surprisingly high off-rates. We find that both factors form dynamic subnuclear hubs, and that Bicoid binding is enriched within Zelda hubs. Remarkably, these hubs are both short lived and interact only transiently with sites of active Bicoid-dependent transcription. Based on our observations, we hypothesize that, beyond simply forming bridges between DNA and the transcription machinery, transcription factors can organize other proteins into hubs that transiently drive multiple activities at their gene targets. Editorial note: This article has been through an editorial process in which the authors decide how to respond to the issues raised during peer review. The Reviewing Editor's assessment is that all the issues have been addressed (see decision letter ).