Uncovering nick DNA binding by LIG1 at the single-molecule level.

DNA ligases repair the strand breaks are made continually and naturally throughout the genome, if left unrepaired and allowed to persist, they can lead to genome instability in the forms of lethal double-strand (ds) breaks, deletions, and duplications. DNA ligase 1 (LIG1) joins Okazaki fragments during the replication machinery and seals nicks at the end of most DNA repair pathways. Yet, how LIG1 recognizes its target substrate is entirely missing. Here, we uncover the dynamics of nick DNA binding by LIG1 at the single-molecule level. Our findings reveal that LIG1 binds to dsDNA both specifically and non-specifically and exhibits diffusive behavior to form a stable complex at the nick. Furthermore, by comparing with the LIG1 C-terminal protein, we demonstrate that the N-terminal non-catalytic region promotes binding enriched at nick sites and facilitates an efficient nick search process by promoting 1D diffusion along the DNA. Our findings provide a novel single-molecule insight into the nick binding by LIG1, which is critical to repair broken phosphodiester bonds in the DNA backbone to maintain genome integrity.

Transcription shapes 3D chromatin organization by interacting with loop extrusion.

Cohesin folds mammalian interphase chromosomes by extruding the chromatin fiber into numerous loops. "Loop extrusion" can be impeded by chromatin-bound factors, such as CTCF, which generates characteristic and functional chromatin organization patterns. It has been proposed that transcription relocalizes or interferes with cohesin and that active promoters are cohesin loading sites. However, the effects of transcription on cohesin have not been reconciled with observations of active extrusion by cohesin. To determine how transcription modulates extrusion, we studied mouse cells in which we could alter cohesin abundance, dynamics, and localization by genetic "knockouts" of the cohesin regulators CTCF and Wapl. Through Hi-C experiments, we discovered intricate, cohesin-dependent contact patterns near active genes. Chromatin organization around active genes exhibited hallmarks of interactions between transcribing RNA polymerases (RNAPs) and extruding cohesins. These observations could be reproduced by polymer simulations in which RNAPs were moving barriers to extrusion that obstructed, slowed, and pushed cohesins. The simulations predicted that preferential loading of cohesin at promoters is inconsistent with our experimental data. Additional ChIP-seq experiments showed that the putative cohesin loader Nipbl is not predominantly enriched at promoters. Therefore, we propose that cohesin is not preferentially loaded at promoters and that the barrier function of RNAP accounts for cohesin accumulation at active promoters. Altogether, we find that RNAP is an extrusion barrier that is not stationary, but rather, translocates and relocalizes cohesin. Loop extrusion and transcription might interact to dynamically generate and maintain gene interactions with regulatory elements and shape functional genomic organization.

Resolving the subtle details of human DNA alkyltransferase lesion search and repair mechanism by single-molecule studies.

SignificanceWe directly visualize DNA translocation and lesion recognition by the O6-alkylguanine DNA alkyltransferase (AGT). Our data show bidirectional movement of AGT monomers and clusters on undamaged DNA that depended on Zn2+ occupancy of AGT. A role of cooperative AGT clusters in enhancing lesion search efficiencies by AGT has previously been proposed. Surprisingly, our data show no enhancement of DNA translocation speed by AGT cluster formation, suggesting that AGT clusters may serve a different role in AGT function. Our data support preferential cluster formation by AGT at alkyl lesions, suggesting a role of these clusters in stabilizing lesion-bound complexes. From our data, we derive a new model for the lesion search and repair mechanism of AGT.

Chromosome organization by one-sided and two-sided loop extrusion.

SMC complexes, such as condensin or cohesin, organize chromatin throughout the cell cycle by a process known as loop extrusion. SMC complexes reel in DNA, extruding and progressively growing DNA loops. Modeling assuming two-sided loop extrusion reproduces key features of chromatin organization across different organisms. In vitro single-molecule experiments confirmed that yeast condensins extrude loops, however, they remain anchored to their loading sites and extrude loops in a 'one-sided' manner. We therefore simulate one-sided loop extrusion to investigate whether 'one-sided' complexes can compact mitotic chromosomes, organize interphase domains, and juxtapose bacterial chromosomal arms, as can be done by 'two-sided' loop extruders. While one-sided loop extrusion cannot reproduce these phenomena, variants can recapitulate in vivo observations. We predict that SMC complexes in vivo constitute effectively two-sided motors or exhibit biased loading and propose relevant experiments. Our work suggests that loop extrusion is a viable general mechanism of chromatin organization.

The different molecules of DNA in a cell are called chromosomes, and they change shape dramatically when cells divide. Ordinarily, chromosomes are packaged by proteins called histones to make thick fibres called chromatin. Chromatin fibres are further folded into a sparse collection of loops. These loops are important not only to make genetic material fit inside a cell, but also to make distant regions of the chromosomes interact with each other, which is important to regulate gene activities. The fibres compact to prepare for cell division: they fold into a much denser series of loops. This is a remarkable physical feat in which tiny protein machines wrangle lengthy strands of DNA. A process called loop extrusion could explain how chromatin folding works. In this process, ring-like protein complexes known as SMC complexes would act as motors that can form loops. SMC complexes could bind a chromatin fibre and reel it in to form the loops, with the density of loops increasing before cell division to further compact the chromosomes. Looping by SMC complexes has been observed in a variety of cell types, including mammalian and bacterial cells. From these studies, loop extrusion is generally assumed to be 'two-sided'. This means that each SMC complex reels in the chromatin on both sides of it, thus growing the chromatin loop. However, imaging individual SMC complexes bound to single molecules of DNA showed that extrusion can be asymmetric, or 'one-sided'. These observations show the SMC complex remains anchored in place and the chromatin is reeled in and extruded by only one side of the complex. So Banigan, van den Berg, Brandão et al. created a computer model to test whether the mechanism of one-sided extrusion could produce chromosomes that are organised, compact, and ready for cell division, like two-sided extrusion can. To answer this question, Banigan, van den Berg, Brandão et al. analysed imaging experiments and data that had been collected using a technique that captures how chromatin fibres are arranged inside cells. This was paired with computer simulations of chromosomes bound by SMC protein complexes. The simulations and analysis found that the simplest one-sided loop extrusion complexes generally cannot reproduce the same patterns of chromatin loops as two-sided complexes. However, a few specific variations of one-sided extrusion can actually recapitulate correct chromatin folding and organisation. These results show that some aspects of chromosome organization can be attained by one-sided extrusion, but many require two-sided extrusion. Banigan, van den Berg, Brandão et al. explain how the simulated mechanisms of loop extrusion could be consistent with seemingly contradictory observations from different sets of experiments. Altogether, they demonstrate that loop extrusion is a viable general mechanism to explain chromatin organisation, and that it likely possesses physical capabilities that have yet to be observed experimentally.

RNA polymerases as moving barriers to condensin loop extrusion.

To separate replicated sister chromatids during mitosis, eukaryotes and prokaryotes have structural maintenance of chromosome (SMC) condensin complexes that were recently shown to organize chromosomes by a process known as DNA loop extrusion. In rapidly dividing bacterial cells, the process of separating sister chromatids occurs concomitantly with ongoing transcription. How transcription interferes with the condensin loop-extrusion process is largely unexplored, but recent experiments have shown that sites of high transcription may directionally affect condensin loop extrusion. We quantitatively investigate different mechanisms of interaction between condensin and elongating RNA polymerases (RNAPs) and find that RNAPs are likely steric barriers that can push and interact with condensins. Supported by chromosome conformation capture and chromatin immunoprecipitation for cells after transcription inhibition and RNAP degradation, we argue that translocating condensins must bypass transcribing RNAPs within ∼1 to 2 s of an encounter at rRNA genes and within ∼10 s at protein-coding genes. Thus, while individual RNAPs have little effect on the progress of loop extrusion, long, highly transcribed operons can significantly impede the extrusion process. Our data and quantitative models further suggest that bacterial condensin loop extrusion occurs by 2 independent, uncoupled motor activities; the motors translocate on DNA in opposing directions and function together to enlarge chromosomal loops, each independently bypassing steric barriers in their path. Our study provides a quantitative link between transcription and 3D genome organization and proposes a mechanism of interactions between SMC complexes and elongating transcription machinery relevant from bacteria to higher eukaryotes.

Crowding-induced transcriptional bursts dictate polymerase and nucleosome density profiles along genes.

During eukaryotic transcription, RNA polymerase (RNAP) translocates along DNA molecules covered with nucleosomes and other DNA binding proteins. Though the interactions between a single nucleosome and RNAP are by now fairly well understood, this understanding has not been synthesized into a description of transcription on crowded genes, where multiple RNAP transcribe through nucleosomes while preserving the nucleosome coverage. We here take a deductive modeling approach to establish the consequences of RNAP-nucleosome interactions for transcription in crowded environments. We show that under physiologically crowded conditions, the interactions of RNAP with nucleosomes induce a strong kinetic attraction between RNAP molecules, causing them to self-organize into stable and moving pelotons. The peloton formation quantitatively explains the observed nucleosome and RNAP depletion close to the initiation site on heavily transcribed genes. Pelotons further translate into short-timescale transcriptional bursts at termination, resulting in burst characteristics consistent with instances of bursty transcription observed in vivo. To facilitate experimental testing of our proposed mechanism, we present several analytic relations that make testable quantitative predictions.

Slow unloading leads to DNA-bound ß2-sliding clamp accumulation in live Escherichia coli cells.

The ubiquitous sliding clamp facilitates processivity of the replicative polymerase and acts as a platform to recruit proteins involved in replication, recombination and repair. While the dynamics of the E. coli ß2-sliding clamp have been characterized in vitro, its in vivo stoichiometry and dynamics remain unclear. To probe both ß2-clamp dynamics and stoichiometry in live E. coli cells, we use custom-built microfluidics in combination with single-molecule fluorescence microscopy and photoactivated fluorescence microscopy. We quantify the recruitment, binding and turnover of ß2-sliding clamps on DNA during replication. These quantitative in vivo results demonstrate that numerous ß2-clamps in E. coli remain on the DNA behind the replication fork for a protracted period of time, allowing them to form a docking platform for other enzymes involved in DNA metabolism.