Taekjip Ha

Chromatin-modifying enzymes must engage their substrate, act on it and release it, but which step limits their turnover is unclear. Using three-color single-molecule fluorescence, we directly visualized the yeast deubiquitinase Ubp10 removing ubiquitin from H2B on individual nucleosomes. Ubp10 cleaved the mark within seconds of binding, without detectable DNA unwrapping, yet it did not let go: ~80% of nucleosomes remained bound after 45 minutes, and unmodified nucleosomes formed similarly long-lived complexes. Ubp10 turnover is therefore limited by release rather than catalysis. The histone chaperone FACT relieved this bottleneck by accelerating Ubp10-nucleosome dissociation. In living yeast, nuclear depletion of the FACT subunit Spt16 increased the slowly diffusing, chromatin-bound fraction of nucleoplasmic Ubp10. These findings identify FACT-dependent release as a likely mechanism for recycling Ubp10 on chromatin. More broadly, a nucleosome can sequester the enzyme acting on it, making enzyme release a step at which chromatin modification can be controlled.

Biomolecular condensates organize essential cellular processes, but how their internal architecture regulates molecular behavior in living cells remains poorly understood. Here, we use MINFLUX nanoscopy for quantitative single-molecule imaging and live-cell tracking to resolve the organization and dynamics of specific macromolecules within nuclear speckles. Rather than behaving as uniform liquid-like compartments, nuclear speckles are hierarchically organized into dynamic nanoscale interaction networks. Proteins and RNAs assemble into molecular clusters that generate transient interaction nanodomains, where individual molecules alternate between diffusion and confinement while exploring higher-order networks. Physiological and genetic perturbations selectively remodel molecular clustering and nanodomain organization, and these changes are not consistently coupled to changes in the abundance of the measured components. Together, these findings reveal a multiscale organization of nuclear speckles in which molecular clustering, transient confinement, and dynamic exchange represent distinct but interconnected features of condensate architecture in living cells.

Molecular chaperones interact with non-native proteins, playing crucial roles in preventing misfolding and in enabling efficient folding in the cellular environment. Trigger factor is a bacterial chaperone that binds to ribosomes, interacting with nascent polypeptides emerging from the ribosome and guiding their early folding steps. In contrast to the central role of the chaperone in promoting folding of newly synthesized proteins, its dynamic interactions with nascent chains emerging from the ribosome remain poorly understood. Here, we use single-molecule fluorescence and optical tweezers approaches to directly observe and characterize trigger factor interactions with a ribosome-bound client protein at increasing chain lengths. We find that trigger factor dwell times on ribosome–nascent chain complexes increase non-monotonically with nascent chain length, and that binding to nascent proteins is best described by a combination of multiple weak, dynamic interactions that are established after the chaperone docks onto the ribosome and evolve during polypeptide elongation. Application of mechanical force perturbs trigger factor binding, supporting a multivalent interaction model. This binding mode may help to stabilize nascent proteins against misfolding while allowing them to dynamically sample conformational space in search of their native structures.

Replicative and accessory helicases are essential for maintaining genome integrity during DNA replication and repair. Rep helicase belongs to superfamily 1 (SF1), the first helicase family solved at atomic resolution and a longstanding model for how helicases couple ATP hydrolysis to movement on DNA. Using optical tweezers combined with fluorescence imaging, we characterize how Rep translocates and navigates diverse nucleic acid and protein obstacles. We directly observe highly processive translocation on ssDNA (∼300 nt/s over >13,000 nt) and show that this directional movement can drive single-stranded DNA-binding proteins (bacterial SSB, archaeal SSB, and eukaryotic RPA) along ssDNA without slowing. On encountering a short duplex barrier (≤60 bp), either a DNA duplex or an RNA/DNA hybrid, Rep frequently bypasses it, with termination and unwinding also observed. Unexpectedly, we identify a mode in which Rep transitions from ssDNA onto dsDNA and translocates directionally along the duplex with high processivity(>12,000bp) and at a much higher speed than on ssDNA (∼1100 bp/s). AlphaFold3 modeling is consistent with Rep tracking a single strand while keeping the duplex intact. To our knowledge, this is the first demonstration of processive dsDNA translocation by an SF1 helicase, and it broadens models for how translocases operate on DNA during replication and repair.

Precise regulation of Cas9 activity is essential to minimize off-target effects, mosaicism, chromosomal alterations, immunogenicity, and genotoxicity in genome editing. Although type II anti-CRISPR proteins (Acrs) can inhibit and regulate Cas9, their size and anionic charge generally prevent them from crossing the cell membrane. Existing Acr delivery methods employing vectors or electroporation are either slow and persistent or require external equipment, limiting their therapeutic utility. To address these challenges, we developed a cell-permeable Acr (6×NLS-Acr), which uses nuclear localization signals (NLSs) to cross the cell membrane. We conjugated 6×NLS-Acr to a fluorescent dye to elucidate its cellular entry mechanism and directly visualized its binding to a fluorescent Cas9·gRNA complex to study its inhibitory mechanism. 6×NLS-Acr (IC50 = 0.47 μM) directly transduces human cells, including immortalized cell lines, embryonic stem cells, and 3D cell cultures, within 5 min, inhibiting up to 99% of Cas9 activity and increasing genome-editing specificity by nearly 100%. We further compared 6×NLS-Acr with our anthrax-derived Acr delivery platform. Our results demonstrate that 6×NLS-Acr is the most efficacious cell-permeable CRISPR-Cas inhibitor, significantly enhancing the precision and therapeutic potential of CRISPR-based genome editing.

Deciphering the spatial organization of molecular machines that copy the genome remains a fundamental challenge in biology. Essential for eukaryotic DNA replication, Mini-Chromosome Maintenance (MCM2-7) helicases are loaded during G1 as double hexamers (DHs) to license replication origins. Upon activation in S phase, each DH is thought to split into two single hexamers (SHs) that form the active CMG helicases and travel bidirectionally. However, the field has long been divided: biochemical and structural studies define CMG helicases as autonomous, independent motors, while genomic and cellular imaging assays suggest sister replisomes remain physically coupled within replication factories. Here, we use MINFLUX nanoscopy to localize individual MCM complexes down to nanometer precision in situ, directly resolving DHs in human cells and capturing their separation into SHs upon origin firing. We find that the resulting sister replisomes do not diffuse apart: they remain coupled at a characteristic distance of ∼40 nm throughout S phase. Depletion experiments identify two distinct contributions to this coupling: local, protein-mediated tethering by the AND1 scaffold, and higher-order spatial confinement dependent on cohesin, which is dispensable for MCM loading in G1 but required to maintain coupling in S phase. By linking the nanometer-scale architecture of the replisome to the genome-wide topology of replication fountains, these findings provide direct spatial evidence that sister forks are coupled during DNA synthesis and define the molecular forces that organize replisomes within their native nuclear context.

Piconewton forces borne by individual proteins within complex assemblies underlie cell adhesion, migration, and tissue morphogenesis, yet remain difficult to image in living systems. Here, we introduce WHaloForce, a chemigenetic HaloTag-based tension sensor that converts force-dependent relief of tryptophan-mediated dye quenching into a fluorescence lifetime change. Optical tweezers revealed a switch-like unquenching transition near 5 pN, and cellular measurements demonstrated dynamic responses to actomyosin activity. WHaloForce enabled quantitative tension imaging of diverse force-bearing proteins (vinculin, E-cadherin, α-catenin, and laminin) in mammalian cells, mouse tissue, and C. elegans. Bright synthetic dyes made tension measurements possible at endogenous expression levels. In C. elegans, vinculin and laminin showed opposite tension patterns between tissues, highlighting tissue-specific force transmission through adhesions and the extracellular matrix. We also identified ovulation as a source of molecular loading within the spermathecal laminin network. WHaloForce thus offers a modular platform for imaging spatiotemporal tension patterns in living systems.

In homologous recombination, DNA repair depends on recombinase filaments finding homologous templates on chromosomes whose topology is continually remodeled by replication and transcription. How dynamic chromosome topology affects homology search and DNA repair remains unknown. By combining live-cell imaging of RecA-mediated homology search with single-molecule assays on topologically-defined DNA substrates, we show that DNA supercoiling acts as a selectivity filter for productive homology search. In Caulobacter crescentus cells, supercoiling is dispensable for the filament movement required for homology search, but required for homology target capture and repair. In vitro , filaments transiently sample both relaxed and supercoiled DNA, yet selectively commit to capture only on negatively supercoiled targets. Mechanistically, we find an extreme kinetic preference (~100-fold) of filaments towards negatively supercoiled DNA targets; this topological preference is independent of DNA compaction, pins plectonemes at the capture site, and is conserved across diverse bacterial RecA homologs. Our findings establish that chromosome topology physically regulates homology search and DNA repair.

Cohesin is a protein complex that shapes 3D genome organization through two distinct mechanisms. First, cohesin tethers replicated chromatids from DNA replication until mitosis. This process, known as sister chromatid cohesion, ensures accurate chromosome segregation and enables high-fidelity DNA repair through homologous recombination between the sister chromatids. Second, cohesin organizes the genome during interphase by dynamically extruding chromatin loops, structures that have key roles in gene regulation. Recent work has shown that, in addition to the well-established repair functions of sister chromatid cohesion, cohesin-mediated chromatin looping is closely linked to the repair of DNA double-strand breaks — one of the most toxic DNA lesions. In this Review, we discuss the central roles of cohesin in maintaining genome stability, with emphasis on the cellular response to DNA double-strand breaks. We review how dynamic loop structures facilitate signalling of repair events and promote long-range chromatin motions that underpin the repair process. Overall, its dual mode of action — cohesion and loop extrusion — positions cohesin as a central regulator of chromatin architecture and genome maintenance.

DNA replication relies on hexameric, ring-shaped helicases to unwind parental DNA for supporting fork progression over tens of thousands of base pairs. Oddly, biochemical studies have suggested that, on their own, replicative helicases are rather limited motors that struggle to couple rapid movement to nucleotide turnover (typically believed to be solely ATP). Here, single-molecule studies reveal that when properly loaded, the Escherichia coli (E. coli) replicative helicase, DnaB, is a tremendously fast single-stranded DNA translocase that moves up to three times more rapidly than the replisome (3 knt/s). Translocation is highly processive, resistant to pulling force/high salt, and can displace short, 3’-tailed DNA duplexes without apparent changes in speed. Surprisingly, we find that the loader for DnaB, DnaC, can use any rNTP or dATP for depositing DnaB onto ssDNA and that the helicase itself also must hydrolyze nucleotide for stable loading. DnaB translocation also turns out to be supported by any r/dNTP, a property shown to extend to the eukaryotic CMG replicative helicase. Overall, the DNA unwinding engines that support cellular replisomes are highly indiscriminate of their fuel source, a feature that may be of utility during times when cells encounter nucleotide pool stress but have committed to DNA synthesis.