Unzipping single DNA molecules to study nucleosome structure and dynamics

Real-Time Multistep Asymmetrical Disassembly of Nucleosomes and Chromatosomes Visualized by High-Speed Atomic Force Microscopy

During replication, expression, and repair of the eukaryotic genome, cellular machinery must access the DNA wrapped around histone proteins forming nucleosomes. These octameric protein·DNA complexes are modular, dynamic, and flexible and unwrap or disassemble either spontaneously or by the action of molecular motors. Thus, the mechanism of formation and regulation of subnucleosomal intermediates has gained attention genome-wide because it controls DNA accessibility. Here, we imaged nucleosomes and their more compacted structure with the linker histone H1 (chromatosomes) using high-speed atomic force microscopy to visualize simultaneously the changes in the DNA and the histone core during their disassembly when deposited on mica. Furthermore, we trained a neural network and developed an automatic algorithm to track molecular structural changes in real time. Our results show that nucleosome disassembly is a sequential process involving asymmetrical stepwise dimer ejection events. The presence of H1 restricts DNA unwrapping, significantly increases the nucleosomal lifetime, and affects the pathway in which heterodimer asymmetrical dissociation occurs. We observe that tetrasomes are resilient to disassembly and that the tetramer core (H3·H4)2 can diffuse along the nucleosome positioning sequence. Tetrasome mobility might be critical to the proper assembly of nucleosomes and can be relevant during nucleosomal transcription, as tetrasomes survive RNA polymerase passage. These findings are relevant to understanding nucleosome intrinsic dynamics and their modification by DNA-processing enzymes.

Conformational Change of Nucleosome Arrays prior to Phase Separation

Chromatin phase transition serves as a regulatory mechanism for eukaryotic transcription. Understanding this process requires the characterization of the nucleosome array structure in response to external stimuli prior to phase separation. However, the intrinsic flexibility and heterogeneity hinders the arrays' structure determination. Here we exploit advances in cryogenic electron tomography (cryo-ET) to determine the three-dimensional (3D) structure of each individual particle of mono-, di-, tri-, and tetranucleosome arrays. Statistical analysis reveals the ionic strength changes the angle between the DNA linker and nucleosome core particle (NCP), which regulate the overall morphology of nucleosome arrays. The finding that one-third of the arrays in the presence of H1 contain an NCP invaded by foreign DNA suggests an alternative function of H1 in constructing nucleosomal networks. The new insights into the nucleosome conformational changes prior to the intermolecular interaction stage extends our understanding of chromatin phase separation regulation.

Polarity of the CRISPR roadblock to transcription

CRISPR (clustered regularly interspaced short palindromic repeats) utility relies on a stable Cas effector complex binding to its target site. However, a Cas complex bound to DNA may be removed by motor proteins carrying out host processes and the mechanism governing this removal remains unclear. Intriguingly, during CRISPR interference, RNA polymerase (RNAP) progression is only fully blocked by a bound endonuclease-deficient Cas (dCas) from the protospacer adjacent motif (PAM)-proximal side. By mapping dCas-DNA interactions at high resolution, we discovered that the collapse of the dCas R-loop allows Escherichia coli RNAP read-through from the PAM-distal side for both Sp-dCas9 and As-dCas12a. This finding is not unique to RNAP and holds for the Mfd translocase. This mechanistic understanding allowed us to modulate the dCas R-loop stability by modifying the guide RNAs. This work highlights the importance of the R-loop in dCas-binding stability and provides valuable mechanistic insights for broad applications of CRISPR technology.

Quantifying epigenetic modulation of nucleosome breathing by high-throughput AFM imaging

Nucleosomes are the basic units of chromatin and critical for storage and expression of eukaryotic genomes. Chromatin accessibility and gene readout are heavily regulated by epigenetic marks, in which post-translational modifications of histones play a key role. However, the mode of action and the structural implications at the single-molecule level of nucleosomes is still poorly understood. Here we apply a high-throughput atomic force microscopy imaging and analysis pipeline to investigate the conformational landscape of the nucleosome variants three additional methyl groups at lysine 36 of histone H3 (H3K36me3), phosphorylation of H3 histones at serine 10 (H3S10phos), and acetylation of H4 histones at lysines 5, 8, 12, and 16 (H4K5/8/12/16ac). Our data set of more than 25,000 nucleosomes reveals nucleosomal unwrapping steps corresponding to 5-bp DNA. We find that H3K36me3 nucleosomes unwrap significantly more than wild-type nucleosomes and additionally unwrap stochastically from both sides, similar to centromere protein A (CENP-A) nucleosomes and in contrast to the highly anticooperative unwrapping of wild-type nucleosomes. Nucleosomes with H3S10phos or H4K5/8/12/16ac modifications show unwrapping populations similar to wild-type nucleosomes and also retain the same level of anticooperativity. Our findings help to put the mode of action of these modifications into context. Although H3K36me3 likely acts partially by directly affecting nucleosome structure on the single-molecule level, H3S10phos and H4K5/8/12/16ac must predominantly act through higher-order processes. Our analysis pipeline is readily applicable to other nucleosome variants and will facilitate future high-resolution studies of the conformational landscape of nucleoprotein complexes.

Resonator nanophotonic standing-wave array trap for single-molecule manipulation and measurement

Nanophotonic tweezers represent emerging platforms with significant potential for parallel manipulation and measurements of single biological molecules on-chip. However, trapping force generation represents a substantial obstacle for their broader utility. Here, we present a resonator nanophotonic standing-wave array trap (resonator-nSWAT) that demonstrates significant force enhancement. This platform integrates a critically-coupled resonator design to the nSWAT and incorporates a novel trap reset scheme. The nSWAT can now perform standard single-molecule experiments, including stretching DNA molecules to measure their force-extension relations, unzipping DNA molecules, and disrupting and mapping protein-DNA interactions. These experiments have realized trapping forces on the order of 20 pN while demonstrating base-pair resolution with measurements performed on multiple molecules in parallel. Thus, the resonator-nSWAT platform now meets the benchmarks of a table-top precision optical trapping instrument in terms of force generation and resolution. This represents the first demonstration of a nanophotonic platform for such single-molecule experiments.

Applications of nanophotonic tweezers have been limited by the low trapping force. Here, the authors present enhanced force generation in a nanophotonic standing-wave array trap by integrating a critically-coupled resonator design and demonstrate common single-molecule experiments.

Angular Optical Trapping to Directly Measure DNA Torsional Mechanics

Angular optical trapping (AOT) is a powerful technique that permits direct angular manipulation of a trapped particle with simultaneous measurement of torque and rotation, while also retaining the capabilities of position and force detection. This technique provides unique approaches to investigate the torsional properties of nucleic acids and DNA-protein complexes, as well as impacts of torsional stress on fundamental biological processes, such as transcription and replication. Here we describe the principle, construction, and calibration of the AOT in detail and provide a guide to the performance of single-molecule torque measurements on DNA molecules. We include the constant-force method and, notably, a new constant-extension method that enables measurement of the twist persistence length of both extended DNA, under an extremely low force, and plectonemic DNA. This chapter can assist in the implementation and application of this technique for general researchers in the single-molecule field.

Effects of size, cooperativity, and competitive binding on protein positioning on DNA

Accurate positioning of proteins on chromosomal DNA is crucial for its proper organization as well as gene transcription regulation. Recent experiments revealed existence of periodic patterns of nucleoprotein complexes on DNA, which frequently cannot be explained by sequence-dependent binding of proteins. Previous theoretical studies suggest that such patterns typically emerge as a result of the proteins' volume-exclusion effect. However, the role of other physical factors in patterns' formation, such as the length of DNA, its sequence heterogeneity, and protein binding cooperativity/binding competition to DNA, remains unclear. To address these less understood yet important aspects, we investigated potential effects of these factors on protein positioning on finite-size DNA by using transfer-matrix calculations. It has been found that upon binding to DNA, proteins form oscillatory patterns that span over the length of up to ∼10 times the size of the protein binding site, with the shape of the patterns being strongly dependent on the length of DNA and the proteins' binding cooperativity to DNA. Furthermore, calculations showed that small variations in the proteins' affinity to DNA due to its sequence heterogeneity do not much change the main geometric characteristics of the observed protein patterns. Finally, competition between two different types of proteins for binding to DNA has been found to lead to formation of highly diverse and complex alternating positioning of the two proteins. Altogether, these results provide new insights into the roles of physicochemical properties of proteins, the DNA length, and DNA-binding competition between proteins in formation of protein positioning patterns on DNA.

Retroviral prototype foamy virus intasome binding to a nucleosome target does not determine integration efficiency

Retroviral integrases must navigate host DNA packaged as chromatin during integration of the viral genome. Prototype foamy virus (PFV) integrase (IN) forms a tetramer bound to two viral DNA (vDNA) ends in a complex termed an intasome. PFV IN consists of four domains: the amino terminal extension domain (NED), amino terminal domain (NTD), catalytic core domain (CCD), and carboxyl terminal domain (CTD). The domains of the two inner IN protomers have been visualized, as well as the CCDs of the two outer IN protomers. However, the roles of the amino and carboxyl terminal domains of the PFV intasome outer subunits during integration to a nucleosome target substrate are not clear. We used the well-characterized 601 nucleosome to assay integration activity as well as intasome binding. PFV intasome integration to 601 nucleosomes occurs in clusters at four independent sites. We find that the outer protomer NED and NTD domains have no significant effects on integration efficiency, site selection, or binding. The CTDs of the outer PFV intasome subunits dramatically affect nucleosome binding but have little effect on total integration efficiency. The outer PFV IN CTDs did significantly alter the integration efficiency at one site. Histone tails also significantly affect intasome binding, but have little impact on PFV integration efficiency or site selection. These results indicate that binding to nucleosomes does not correlate with integration efficiency and suggests most intasome-binding events are unproductive.

High-throughput AFM analysis reveals unwrapping pathways of H3 and CENP-A nucleosomes

Nucleosomes, the fundamental units of chromatin, regulate readout and expression of eukaryotic genomes. Single-molecule experiments have revealed force-induced nucleosome accessibility, but a high-resolution unwrapping landscape in the absence of external forces is currently lacking. Here, we introduce a high-throughput pipeline for the analysis of nucleosome conformations based on atomic force microscopy and automated, multi-parameter image analysis. Our data set of ∼10 000 nucleosomes reveals multiple unwrapping states corresponding to steps of 5 bp DNA. For canonical H3 nucleosomes, we observe that dissociation from one side impedes unwrapping from the other side, but in contrast to force-induced unwrapping, we find only a weak sequence-dependent asymmetry. Notably, centromeric CENP-A nucleosomes do not unwrap anti-cooperatively, in stark contrast to H3 nucleosomes. Finally, our results reconcile previous conflicting findings about the differences in height between H3 and CENP-A nucleosomes. We expect our approach to enable critical insights into epigenetic regulation of nucleosome structure and stability and to facilitate future high-throughput AFM studies that involve heterogeneous nucleoprotein complexes.

Observing protein interaction dynamics to chemically defined chromatin fibers by colocalization single-molecule fluorescence microscopy

In eukaryotic cells, the genome is packaged into chromatin and exists in different states, ranging from open euchromatic regions to highly condensed heterochromatic regions. Chromatin states are highly dynamic and are organized by an interplay of histone post-translational modifications and effector proteins, both of which are central in the regulation of gene expression. For this, chromatin effector proteins must first search the nucleus for their targets, before binding and performing their role. A key question is how chromatin effector proteins search, interact with and alter the different chromatin environments. Here we present a modular fluorescence based in vitro workflow to directly observe dynamic interactions of effector proteins with defined chromatin fibres, replicating different chromatin states. We discuss the design and creation of chromatin assemblies, the synthesis of modified histones, the fabrication of microchannels and the approach to data acquisition and analysis. All of this with the aim to better understand the complex in vivo relationship between chromatin structure and gene expression.

Constructing arrays of nucleosome positioning sequences using Gibson Assembly for single-molecule studies

As the basic building blocks of chromatin, nucleosomes play a key role in dictating the accessibility of the eukaryotic genome. Consequently, nucleosomes are involved in essential genomic transactions such as DNA transcription, replication and repair. In order to unravel the mechanisms by which nucleosomes can influence, or be altered by, DNA-binding proteins, single-molecule techniques are increasingly employed. To this end, DNA molecules containing a defined series of nucleosome positioning sequences are often used to reconstitute arrays of nucleosomes in vitro. Here, we describe a novel method to prepare DNA molecules containing defined arrays of the ‘601’ nucleosome positioning sequence by exploiting Gibson Assembly cloning. The approaches presented here provide a more accessible and efficient means to generate arrays of nucleosome positioning motifs, and facilitate a high degree of control over the linker sequences between these motifs. Nucleosomes reconstituted on such arrays are ideal for interrogation with single-molecule techniques. To demonstrate this, we use dual-trap optical tweezers, in combination with fluorescence microscopy, to monitor nucleosome unwrapping and histone localisation as a function of tension. We reveal that, although nucleosomes unwrap at ~20 pN, histones (at least histone H3) remain bound to the DNA, even at tensions beyond 60 pN.

Nucleosome DNA unwrapping does not affect prototype foamy virus integration efficiency or site selection

Eukaryotic DNA binding proteins must access genomic DNA that is packaged into chromatin in vivo. During a productive infection, retroviral integrases (IN) must similarly interact with chromatin to integrate the viral cDNA genome. Here we examine the role of nucleosome DNA unwrapping in the retroviral integrase search for a target site. These studies utilized PFV intasomes that are comprised of a tetramer of PFV IN with two oligomers mimicking the viral cDNA ends. Modified recombinant human histones were used to generate nucleosomes with increased unwrapping rates at different DNA regions. These modifications included the acetylmimetic H3(K56Q) and the chemically engineered H4(K77ac, K79ac). While transcription factors and DNA damage sensors may search nucleosome bound DNA during transient unwrapping, PFV intasome mediated integration appears to be unaffected by increased nucleosome unwrapping. These studies suggest PFV intasomes do not utilize nucleosome unwrapping to search nucleosome targets.

Histone Acetylation Inhibits RSC and Stabilizes the +1 Nucleosome

The +1 nucleosome of yeast genes, within which reside transcription start sites, is characterized by histone acetylation, by the displacement of an H2A-H2B dimer, and by a persistent association with the RSC chromatin-remodeling complex. Here we demonstrate the interrelationship of these characteristics and the conversion of a nucleosome to the +1 state in vitro. Contrary to expectation, acetylation performs an inhibitory role, preventing the removal of a nucleosome by RSC. Inhibition is due to both enhanced RSC-histone interaction and diminished histone-chaperone interaction. Acetylation does not prevent all RSC activity, because stably bound RSC removes an H2A-H2B dimer on a timescale of seconds in an irreversible manner.

Local DNA Sequence Controls Asymmetry of DNA Unwrapping from Nucleosome Core Particles

DNA is tightly wrapped around histone proteins in nucleosome core particles (NCPs) yet must become accessible for processing in the cell. This accessibility, a key component of transcription regulation, is influenced by the properties of both the histone proteins and the DNA itself. Small angle x-ray scattering with contrast variation is used to examine how sequence variations affect DNA unwrapping from NCPs at different salt concentrations. Salt destabilizes NCPs, populating multiple unwrapped states as many possible unwrapping pathways are explored by the complexes. We apply coarse-grained Monte Carlo methods to generate realistic sequence-dependent unwrapped structures for the nucleosomal DNA with thermal variations. An ensemble optimization method is employed to determine the composition of the overall ensemble as electrostatic interactions are weakened. Interesting DNA-sequence-dependent differences are revealed in the unwrapping paths and equilibrium constants. These differences are correlated with specific features within the nucleic acid sequences.

Transcription and Remodeling Produce Asymmetrically Unwrapped Nucleosomal Intermediates

Nucleosomes are disrupted during transcription and other active processes, but the structural intermediates during nucleosome disruption in vivo are unknown. To identify intermediates, we mapped subnucleosomal protections in Drosophila cells using Micrococcal Nuclease followed by sequencing. At the first nucleosome position downstream of the transcription start site, we identified unwrapped intermediates, including hexasomes that lack either proximal or distal contacts. Inhibiting topoisomerases or depleting histone chaperones increased unwrapping, whereas inhibiting release of paused RNAPII or reducing RNAPII elongation decreased unwrapping. Our results indicate that positive torsion generated by elongating RNAPII causes transient loss of histone-DNA contacts. Using this mapping approach, we found that nucleosomes flanking human CTCF insulation sites are similarly disrupted. We also identified diagnostic subnucleosomal particle remnants in cell-free human DNA data as a relic of transcribed genes from apoptosing cells. Thus identification of subnucleosomal fragments from nuclease protection data represents a general strategy for structural epigenomics.

Mfd Dynamically Regulates Transcription via a Release and Catch-Up Mechanism

The bacterial Mfd ATPase is increasingly recognized as a general transcription factor that participates in the resolution of transcription conflicts with other processes/roadblocks. This function stems from Mfd's ability to preferentially act on stalled RNA polymerases (RNAPs). However, the mechanism underlying this preference and the subsequent coordination between Mfd and RNAP have remained elusive. Here, using a novel real-time translocase assay, we unexpectedly discovered that Mfd translocates autonomously on DNA. The speed and processivity of Mfd dictate a "release and catch-up" mechanism to efficiently patrol DNA for frequently stalled RNAPs. Furthermore, we showed that Mfd prevents RNAP backtracking or rescues a severely backtracked RNAP, allowing RNAP to overcome stronger obstacles. However, if an obstacle's resistance is excessive, Mfd dissociates the RNAP, clearing the DNA for other processes. These findings demonstrate a remarkably delicate coordination between Mfd and RNAP, allowing efficient targeting and recycling of Mfd and expedient conflict resolution.

DNA looping mediates nucleosome transfer

Proper cell function requires preservation of the spatial organization of chromatin modifications. Maintenance of this epigenetic landscape necessitates the transfer of parental nucleosomes to newly replicated DNA, a process that is stringently regulated and intrinsically linked to replication fork dynamics. This creates a formidable setting from which to isolate the central mechanism of transfer. Here we utilized a minimal experimental system to track the fate of a single nucleosome following its displacement, and examined whether DNA mechanics itself, in the absence of any chaperones or assembly factors, may serve as a platform for the transfer process. We found that the nucleosome is passively transferred to available dsDNA as predicted by a simple physical model of DNA loop formation. These results demonstrate a fundamental role for DNA mechanics in mediating nucleosome transfer and preserving epigenetic integrity during replication.

Single-Molecule Optical-Trapping Techniques to Study Molecular Mechanisms of a Replisome

The replisome is a multiprotein molecular machinery responsible for the replication of DNA. It is composed of several specialized proteins each with dedicated enzymatic activities, and in particular, helicase unwinds double-stranded DNA and DNA polymerase catalyzes the synthesis of DNA. Understanding how a replisome functions in the process of DNA replication requires methods to dissect the mechanisms of individual proteins and of multiproteins acting in concert. Single-molecule optical-trapping techniques have proved to be a powerful approach, offering the unique ability to observe and manipulate biomolecules at the single-molecule level and providing insights into the mechanisms of molecular motors and their interactions and coordination in a complex. Here, we describe a practical guide to applying these techniques to study the dynamics of individual proteins in the bacteriophage T7 replisome, as well as the coordination among them. We also summarize major findings from these studies, including nucleotide-specific helicase slippage and new lesion bypass pathway in T7 replication.

Protein-DNA and ion-DNA interactions revealed through contrast variation SAXS

Understanding how DNA carries out its biological roles requires knowledge of its interactions with biological partners. Since DNA is a polyanionic polymer, electrostatic interactions contribute significantly. These interactions are mediated by positively charged protein residues or charge compensating cations. Direct detection of these partners and/or their effect on DNA conformation poses challenges, especially for monitoring conformational dynamics in real time. Small-angle x-ray scattering (SAXS) is uniquely sensitive to both the conformation and local environment (i.e. protein partner and associated ions) of the DNA. The primary challenge of studying multi-component systems with SAXS lies in resolving how each component contributes to the measured scattering. Here, we review two contrast variation (CV) strategies that enable targeted studies of the structures of DNA or its associated partners. First, solution contrast variation enables measurement of DNA conformation within a protein-DNA complex by masking out the protein contribution to the scattering profile. We review a specific example, in which the real-time unwrapping of DNA from a nucleosome core particle is measured during salt-induced disassembly. The second method, heavy atom isomorphous replacement, reports the spatial distribution of the cation cloud around duplex DNA by exploiting changes in the scattering strength of cations with varying atomic numbers. We demonstrate the application of this approach to provide the spatial distribution of monovalent cations (Na+, K+, Rb+, Cs+) around a standard 25-base pair DNA. The CV strategies presented here are valuable tools for understanding DNA interactions with its biological partners.

Arginine-phosphate salt bridges between histones and DNA: intermolecular actuators that control nucleosome architecture

Structural bioinformatics and van der Waals density functional theory are combined to investigate the mechanochemical impact of a major class of histone-DNA interactions, namely, the formation of salt bridges between arginine residues in histones and phosphate groups on the DNA backbone. Principal component analysis reveals that the configurational fluctuations of the sugar-phosphate backbone display sequence-specific directionality and variability, and clustering of nucleosome crystal structures identifies two major salt-bridge configurations: a monodentate form in which the arginine end-group guanidinium only forms one hydrogen bond with the phosphate, and a bidentate form in which it forms two. Density functional theory calculations highlight that the combination of sequence, denticity, and salt-bridge positioning enables the histones to apply a tunable mechanochemical stress to the DNA via precise and specific activation of backbone deformations. The results suggest that selection for specific placements of van der Waals contacts, with high-precision control of the spatial distribution of intermolecular forces, may serve as an underlying evolutionary design principle for the structure and function of nucleosomes, a conjecture that is corroborated by previous experimental studies.

Dynamic regulation of transcription factors by nucleosome remodeling

The chromatin landscape and promoter architecture are dominated by the interplay of nucleosome and transcription factor (TF) binding to crucial DNA sequence elements. However, it remains unclear whether nucleosomes mobilized by chromatin remodelers can influence TFs that are already present on the DNA template. In this study, we investigated the interplay between nucleosome remodeling, by either yeast ISW1a or SWI/SNF, and a bound TF. We found that a TF serves as a major barrier to ISW1a remodeling, and acts as a boundary for nucleosome repositioning. In contrast, SWI/SNF was able to slide a nucleosome past a TF, with concurrent eviction of the TF from the DNA, and the TF did not significantly impact the nucleosome positioning. Our results provide direct evidence for a novel mechanism for both nucleosome positioning regulation by bound TFs and TF regulation via dynamic repositioning of nucleosomes.

DOI:http://dx.doi.org/10.7554/eLife.06249.001

Cells contain thousands of genes that are encoded by molecules of DNA. In yeast and other eukaryotic organisms, this DNA is wrapped around proteins called histones to make structures called nucleosomes. This compacts the DNA and allows it to fit inside the tiny nucleus within the cell. The positioning of the nucleosomes influences how tightly packed the DNA is, which in turn influences the activity of genes. Less active genes tend to be found within regions of DNA that are tightly packed, while more active genes are found in less tightly packed regions.

To activate a gene, proteins called transcription factors bind to a section of DNA within the gene called the promoter. Enzymes known as ‘chromatin remodelers’ can alter the locations of nucleosomes on DNA to allow the transcription factors access to the promoters of particular genes. In yeast, the SWI/SNF family of chromatin remodelers can disassemble nucleosomes to promote gene activity, while the ISW1 family organises nucleosomes into closely spaced groups to repress gene activity. However, it is not clear if, or how, chromatin remodelers can influence transcription factors that are already bound to DNA.

Here, Li et al. studied the interactions between a transcription factor and the chromatin remodelers in yeast. The experiment used a piece of DNA that contained a bound transcription factor and a single nucleosome. Li et al. used a technique called ‘single molecule DNA unzipping’, which enabled them to precisely locate the position of the nucleosome and transcription factor before and after the nucleosome was remodeled. The experiments found that a chromatin remodeler called ISW1a moved the nucleosome away from the transcription factor, while a SWI/SNF chromatin remodeler moved the nucleosome towards it.

Significantly, Li et al. also found that a transcription factor is a major barrier to ISW1a's remodeling activity, suggesting that ISW1a may use transcription factors as reference points to position nucleosomes. In contrast, SWI/SNF was able to slide a nucleosome past the transcription factor, which led to the transcription factor falling off the DNA. Therefore, SWI/SNF is able to move transcription factors out of the way to deactivate genes.

Li et al. propose a new model for how chromatin remodelers can move nucleosomes and regulate transcription factors to alter gene activity. A future challenge will be to observe these types of activities in living cells.

DOI:http://dx.doi.org/10.7554/eLife.06249.002

Torque modulates nucleosome stability and facilitates H2A/H2B dimer loss

The nucleosome, the fundamental packing unit of chromatin, has a distinct chirality: 147 bp of DNA are wrapped around the core histones in a left-handed, negative superhelix. It has been suggested that this chirality has functional significance, particularly in the context of the cellular processes that generate DNA supercoiling, such as transcription and replication. However, the impact of torsion on nucleosome structure and stability is largely unknown. Here we perform a detailed investigation of single nucleosome behavior on the high affinity 601 positioning sequence under tension and torque using the angular optical trapping technique. We find that torque has only a moderate effect on nucleosome unwrapping. In contrast, we observe a dramatic loss of H2A/H2B dimers upon nucleosome disruption under positive torque, while (H3/H4)₂ tetramers are efficiently retained irrespective of torsion. These data indicate that torque could regulate histone exchange during transcription and replication.