Topoisomerase enzymes are essential for the regulation of DNA topology. Human topoisomerase IIIα is a Type 1A topoisomerase that exists as a complex with RMI1 and RMI2, known as TRR. The TRR complex can unlink entwined DNA strands and is known to be important for resolving DNA replication and recombination intermediates. It has recently been proposed that TRR can also relax transient negatively supercoiled loops of DNA generated by the translocase PICH and that this activity may help to facilitate the resolution of ultrafine anaphase bridges (UFBs) between segregating sister chromatids. However, the mechanism by which TRR interacts with, and processes, negatively supercoiled DNA is not well understood. Here, we establish a single-molecule strategy to simultaneously measure real-time changes in supercoiling density and visualize the interactions of TRR with underwound DNA using a combination of optical tweezers and fluorescence imaging. We demonstrate that TRR relaxes highly negatively supercoiled DNA in a processive manner and that the timescale for relaxation is less than the expected lifetime of the negatively supercoiled loops generated by PICH. We also show that in the absence of free protein in solution, TRR remains bound to the DNA for long time periods after the torsional stress has been released. Our findings provide a mechanistic basis for how TRR can relax negative supercoils, consistent with its proposed role in UFB resolution. Moreover, our assay could also be widely applied to study the interactions of other families of topoisomerases with negatively supercoiled DNA.

ABSTRACT Human CCCTC-binding factor (CTCF) is a crucial factor in genome organization, regulating chromatin looping and gene expression. The loop extrusion model (LEM) designates cohesin as the sole active player and limits CTCF to a passive barrier, though emerging evidence suggests a more dynamic role. Using complementary single-molecule techniques, combining dual- and quad-trap optical tweezers, fluorescence microscopy, and atomic force microscopy, we show CTCF forms stable, dynamic DNA bridges. We demonstrate CTCF binds with high affinity and cooperativity, undergoes 1D diffusion, and stiffens DNA. Strikingly, CTCF stabilizes DNA loops without cohesin and forms bridges that slide at low force yet resist rupture at forces exceeding dsDNA stability. These findings suggest a modified LEM in which CTCF directly forms dynamic, force-resistant bridges to stabilize cohesin-established loops, revealing an active role in genome organization.

Faithful chromosome segregation during mitosis relies on the formation of compact, individualized chromosomes that withstand drag and spindle-generated forces. Structural failure of mitotic chromosomes under force can disrupt the distribution of genetic material to daughter cells, causing aneuploidy or cancer. The overall mechanical properties of mitotic chromosomes have been suggested to arise from their structural heterogeneity. The magnitude and scale of this heterogeneity have not been measured, leaving its impact on chromosome mechanics unresolved. Here we show that chromosomes are highly mechanically heterogeneous: within one chromosome, the local stiffness can vary by up to two orders of magnitude. This extreme mechanical heterogeneity is exemplified by the centromere, which is an order of magnitude softer than the whole chromosome. These results demonstrate how the mechanical complexity of mitotic chromosomes gives rise to their emergent nonlinear mechanical behaviour, distinct from the polymer properties of their constituents. More broadly, we discuss how structural heterogeneity can shape the nonlinear responses of composite materials, with implications for both understanding biological assemblies and designing new synthetic materials. Analysis of the mechanical properties of mitotic chromosomes is key for understanding the robustness of chromosomes during cell division. It is now shown that chromosomes are highly mechanically heterogeneous.

Abstract Transcription is a vital cellular process in which RNA polymerase produces messenger RNAs (mRNA) from a DNA template. Many transcription systems have been developed to reproduce this process in vitro and in the confined environment of giant unilamellar vesicles (GUVs). However, these systems and the studies employing them often use DNA concentrations significantly higher than those found in natural cells, which typically contain a single DNA copy. In this work, we introduce single-molecule in vitro transcription (smIVT) that overcomes this limitation, enabling the visualization and tracking of individual mRNA molecules both in solution and within GUVs. We achieved this by employing a molecular beacon—a quenched RNA or DNA hairpin probe that becomes fluorescent after binding to the transcript mRNA—and template DNA encoding 32 repeats of the molecular beacon binding sequence. With these, we ensured a signal-to-noise ratio sufficient to detect single mRNAs. We use smIVT to compare the performance of various commercially-available in vitro transcription kits, and to quantitatively describe single molecule transcription inside GUVs. We establish smIVT as a remarkable technique for scrutinizing in vitro transcription with single-molecule resolution.

A quantitative description of nuclear mechanics is crucial for understanding its role in force sensing within eukaryotic cells. Recent studies indicate that the chromatin within the nucleus cannot be treated as a homogeneous material. To elucidate its material properties, we combine optical tweezers manipulation of isolated nuclei with multi-color fluorescence imaging of lamin and chromatin to map the response of nuclei to local deformations. Force spectroscopy reveals nuclear strain stiffening and an exponential force dependence, well described by a hierarchical chain model. Simultaneously, fluorescence data show a higher compliance of chromatin compared to the nuclear envelope at strains <30%. Micrococcal nuclease (MNase) digestion of chromatin results in nuclear softening and can be captured by our model. Additionally, we observe stretching responses showing a lipid tether signature, suggesting that these tethers originate from the nuclear membrane. Our combined approach allows us to elucidate the nuclear force response while mapping the deformation of lamin, (eu)chromatin, and membrane.

RecQ helicases are molecular motor proteins that play an important role in maintaining chromosomal stability by resolving aberrant structures such as Holliday junctions, D-loop, G-quadruplex and hairpins during DNA replication. Bloom syndrome is a disorder caused by mutations in the RecQ helicase BLM that is characterized by a predisposition to multiple types of cancer. BLM is a single-stranded binding protein that unwinds DNA by utilizing the energy released by ATP hydrolysis. Previously, biochemical studies have attempted to study the kinetic step size, with which BLM unwinds DNA, and describe its mechanism of action.

In the context of soft matter and cellular mechanics, microrheology - the use of micron-sized particles to probe the frequency-dependent viscoelastic response of materials – is widely used to shed light onto the mechanics and dynamics of molecular structures. Here we present the implementation of active microrheology in an Acoustic Force Spectroscopy setup (AFMR), which combines multiplexing with the possibility of probing a wide range of forces ( ~ pN to ~nN) and frequencies (0.01–100 Hz). To demonstrate the potential of this approach, we perform active microrheology on biological samples of increasing complexity and stiffness: collagen gels, red blood cells (RBCs), and human fibroblasts, spanning a viscoelastic modulus range of five orders of magnitude. We show that AFMR can successfully quantify viscoelastic properties by probing many beads with high single-particle precision and reproducibility. Finally, we demonstrate that AFMR to map local sample heterogeneities as well as detect cellular responses to drugs.

Transcription is a vital cellular process in which RNA polymerase produces messenger RNAs (mRNA) from a DNA template. On its trajectory toward a viable artificial cell, synthetic biology aims to reliably reproduce transcription in a confined environment and thoroughly understand how it works. To this end, a number of commercially-available/lab-made transcription systems have been established for in vitro applications. Moreover, using fluorescent probes, researchers could follow the changes in RNA concentration over time inside giant unilamellar vesicles (GUVs).

Bacteria, like eukaryotes, use various mechanisms to compact their chromosomal DNA. Unraveling the intricate organization of the bacterial genome within the nucleoid structure requires a full understanding of the vital proteins involved in its formation, the nucleoid-associated proteins (NAPs). NAPs are key players in nucleoid organization, DNA-based processes, and response to physicochemical cues. Classification of NAPs studied across bacterial species according to some of the most well-studied NAPs such as H-NS relies on shared functionality, structural resemblance, and, at times, sequence similarity.

Bacteria densely organize their genome in the form of a nucleoid stored in the cytoplasm. This compact, functional form of DNA is achieved by the use of nucleoid-associated proteins (NAPs), proteins that play a role in DNA organization as well as regulating gene expression. The various NAPs found across bacterial species are classed based on shared functions with well-studied NAPs (H-NS, HU, IHF and Fis). In Bacillus subtilis, the NAP Rok shares functional similarities with H-NS and is proposed to be a H-NS-like protein, however, some fundamental differences in binding activity and response to physio-chemical changes have been observed.

In anaphase, any unresolved DNA entanglements between the segregating sister chromatids can give rise to chromatin bridges. To prevent genome instability, chromatin bridges must be resolved prior to cytokinesis. The SNF2 protein PICH has been proposed to play a direct role in this process through the remodeling of nucleosomes. However, direct evidence of nucleosome remodeling by PICH has remained elusive. Here, we present an in vitro single-molecule assay that mimicks chromatin under tension, as is found in anaphase chromatin bridges.

The cytoskeleton is a network of interlinked filaments inside the cell. It plays a crucial role in maintaining the cellular shape and drives vital processes such as cell division and motility. An array of sophisticated methods exists at the scientists' disposal (such as atomic force microscopy, micropipette aspiration and optical tweezers) to exert forces on cells and investigate their mechanics. However, these methods come with fundamental drawbacks. Firstly, the necessity of direct contact between the sample and the probe potentially introduces artifacts. Secondly, they are often limited to a single cell per measurement. It would therefore be desirable to have a robust and fast technique to characterize the biomechanics of the cytoskeleton inside living cells.

Studying cellular mechanics allows important insights into its cytoskeletal composition, developmental stage, and health. While many force spectroscopy assays exist that allow probing of mechanics of bioparticles, most of them require immobilization of and direct contact with the particle and can only measure a single particle at a time. Here, we introduce quantitative acoustophoresis (QAP) as a simple alternative that uses an acoustic standing wave field to directly determine cellular compressibility and density of many cells simultaneously in a contact-free manner. First, using polymeric spheres of different sizes and materials, we verify that our assay data follow the standard acoustic theory with great accuracy. We furthermore verify that our technique not only is able to measure compressibilities of living cells but can also sense an artificial cytoskeleton inside a biomimetic vesicle. We finally provide a thorough discussion about the expected accuracy our approach provides. To conclude, we show that compared to existing methods, our QAP assay provides a simple yet powerful alternative to study the mechanics of biological and biomimetic particles.

In anaphase, any unresolved DNA entanglements between the segregating sister chromatids can give rise to chromatin bridges. To prevent genome instability, chromatin bridges must be resolved prior to cytokinesis. The SNF2 protein PICH has been proposed to play a direct role in this process through the remodeling of nucleosomes. However, direct evidence of nucleosome remodeling by PICH has remained elusive. Here, we present an in vitro single-molecule assay that mimics chromatin under tension, as is found in anaphase chromatin bridges. Applying a combination of dual-trap optical tweezers and fluorescence imaging of PICH and histones bound to a nucleosome-array construct, we show that PICH is a tension- and ATP-dependent nucleosome remodeler that facilitates nucleosome unwrapping and then subsequently slides remaining histones along the DNA. This work elucidates the role of PICH in chromatin-bridge dissolution, and might provide molecular insights into the mechanisms of related SNF2 proteins.

DNA structural transitions facilitate genomic processes, mediate drug-DNA interactions, and inform the development of emerging DNA-based biotechnology. Here, we use concurrent fluorescence polarization imaging and DNA manipulation experiments to probe the structure of DNA conformations that form under high tension and topological constraints. First, we employ polarization imaging of DNA intercalators (small dye molecules that slide between the DNA base-pairs) to probe the structure of S-DNA, an elongated conformation that can be accessed by mechanical overstretching of (relaxed) B-DNA.

Self-assembling patchy colloidal particles form a promising platform to create designer soft materials. To dress such systems with mechanical functionality, one can take inspiration from biological structures such as the cell's cytoskeleton, which consists of semiflexible filaments, whose mechanical behavior give the cell its unique mechanical properties. Here we present mechanical experiments on analogs of biological fibers, semiflexible ``colloidal polymers'' made from bonded patchy colloidal particles. We use optical tweezers to probe their extreme mechanics under increasingly high compressions and we reveal a rich nonlinear mechanical response involving buckling, viscoelastic creep, and ultimately break-up. We characterize and model this response using elastic and viscoelastic models involving Euler buckling and stress relaxation. This allows us to identify the critical Euler buckling force, and relate the critical bending at break-up to the finite patch size of the colloids. These results demonstrate the crucial role of the patch-patch interactions in the mechanics of self-assembled colloidal materials, and they provide mechanical relationships that are essential to design functional colloidal architectures inspired by nature.

Topoisomerase IIIα is a type 1A topoisomerase that forms a complex with RMI1 and RMI2 called TRR in human cells. TRR plays an essential role in resolving DNA replication and recombination intermediates, often alongside the helicase BLM. While the TRR catalytic cycle is known to involve a protein-mediated single-stranded (ss)DNA gate, the detailed mechanism is not fully understood. Here, we probe the catalytic steps of TRR using optical tweezers and fluorescence microscopy. We demonstrate that TRR forms an open gate in ssDNA of 8.5 ± 3.8 nm, and directly visualize binding of a second ssDNA or double-stranded (ds)DNA molecule to the open TRR-ssDNA gate, followed by catenation in each case. Strikingly, dsDNA binding increases the gate size (by ~16%), while BLM alters the mechanical flexibility of the gate. These findings reveal an unexpected plasticity of the TRR-ssDNA gate size and suggest that TRR-mediated transfer of dsDNA may be more relevant in vivo than previously believed.

Topoisomerases are essential enzymes that regulate DNA topology. Type 1A family topoisomerases are found in nearly all living organisms and are unique in that they require single-stranded (ss)DNA for activity. These enzymes are vital for maintaining supercoiling homeostasis and resolving DNA entanglements generated during DNA replication and repair. While the catalytic cycle of Type 1A topoisomerases has been long-known to involve an enzyme-bridged ssDNA gate that allows strand passage, a deeper mechanistic understanding of these enzymes has only recently begun to emerge. This knowledge has been greatly enhanced through the combination of biochemical studies and increasingly sophisticated single-molecule assays based on magnetic tweezers, optical tweezers, atomic force microscopy and Förster resonance energy transfer. In this review, we discuss how single-molecule assays have advanced our understanding of the gate opening dynamics and strand-passage mechanisms of Type 1A topoisomerases, as well as the interplay of Type 1A topoisomerases with partner proteins, such as RecQ-family helicases. We also highlight how these assays have shed new light on the likely functional roles of Type 1A topoisomerases in vivo and discuss recent developments in single-molecule technologies that could be applied to further enhance our understanding of these essential enzymes.

The combination of DNA force spectroscopy and polarization microscopy of fluorescent DNA intercalator dyes can provide valuable insights into the structure of DNA under tension. These techniques have previously been used to characterize S-DNA-an elongated DNA conformation that forms when DNA overstretches at forces ≥ 65 pN. In this way, it was deduced that the base pairs of S-DNA are highly inclined, relative to those in relaxed (B-form) DNA. However, it is unclear whether and how topological constraints on the DNA may influence the base-pair inclinations under tension. Here, we apply polarization microscopy to investigate the impact of DNA pulling geometry, torsional constraint, and negative supercoiling on the orientations of intercalated dyes during overstretching. In contrast to earlier predictions, the pulling geometry (namely, whether the DNA molecule is stretched via opposite strands or the same strand) is found to have little influence. However, torsional constraint leads to a substantial reduction in intercalator tilting in overstretched DNA, particularly in AT-rich sequences. Surprisingly, the extent of intercalator tilting is similarly reduced when the DNA molecule is negatively supercoiled up to a critical supercoiling density (corresponding to ∼70% reduction in the linking number). We attribute these observations to the presence of P-DNA (an overwound DNA conformation). Our results suggest that intercalated DNA preferentially flanks regions of P-DNA rather than those of S-DNA and also substantiate previous suggestions that P-DNA forms predominantly in AT-rich sequences.

Abstract Fluorescence microscopy is invaluable to a range of biomolecular analysis approaches. The required labeling of proteins of interest, however, can be challenging and potentially perturb biomolecular functionality as well as cause imaging artefacts and photo bleaching issues. Here, we introduce inverse (super-resolution) imaging of unlabeled proteins bound to DNA. In this new method, we use DNA-binding fluorophores that transiently label bare DNA but not protein-bound DNA. In addition to demonstrating diffraction-limited inverse imaging, we show that inverse Binding-Activated Localization Microscopy or ‘iBALM’ can resolve biomolecular features smaller than the diffraction limit. The current detection limit is estimated to lie at features between 5 and 15 nm in size. Although the current image-acquisition times preclude super-resolving fast dynamics, we show that diffraction-limited inverse imaging can reveal molecular mobility at ∼0.2 s temporal resolution and that the method works both with DNA-intercalating and non-intercalating dyes. Our experiments show that such inverse imaging approaches are valuable additions to the single-molecule toolkit that relieve potential limitations posed by labeling.