Single-molecule imaging of DNA curtains reveals intrinsic energy landscapes for nucleosome deposition

Studying Structure and Functions of Nucleosomes with Atomic Force Microscopy

Chromatin is an epigenetic platform for implementation of DNA-dependent processes. Nucleosome, as a basic level of chromatin compaction, largely determines its properties and structure. In the study of nucleosomes structure and functions physicochemical tools are actively used, such as magnetic and optical "tweezers", "DNA curtains", nuclear magnetic resonance, X-ray crystallography, and cryogenic electron microscopy, as well as optical methods based on Förster resonance energy transfer. Despite the fact that these approaches make it possible to determine a wide range of structural and functional characteristics of chromatin and nucleosomes with high spatial and time resolution, atomic force microscopy (AFM) complements the capabilities of these methods. The results of structural studies of nucleosome focusing on the AFM method development are presented in this review. The possibilities of AFM are considered in the context of application of other physicochemical approaches.

Dynamic 1D search and processive nucleosome translocations by RSC and ISW2 chromatin remodelers

Eukaryotic gene expression is linked to chromatin structure and nucleosome positioning by ATP-dependent chromatin remodelers that establish and maintain nucleosome-depleted regions (NDRs) near transcription start sites. Conserved yeast RSC and ISW2 remodelers exert antagonistic effects on nucleosomes flanking NDRs, but the temporal dynamics of remodeler search, engagement, and directional nucleosome mobilization for promoter accessibility are unknown. Using optical tweezers and two-color single-particle imaging, we investigated the Brownian diffusion of RSC and ISW2 on free DNA and sparse nucleosome arrays. RSC and ISW2 rapidly scan DNA by one-dimensional hopping and sliding, respectively, with dynamic collisions between remodelers followed by recoil or apparent co-diffusion. Static nucleosomes block remodeler diffusion resulting in remodeler recoil or sequestration. Remarkably, both RSC and ISW2 use ATP hydrolysis to translocate mono-nucleosomes processively at ~30 bp/s on extended linear DNA under tension. Processivity and opposing push-pull directionalities of nucleosome translocation shown by RSC and ISW2 shape the distinctive landscape of promoter chromatin.

Magnetic tweezers principles and promises

Magnetic tweezers have become popular with the outbreak of single molecule micromanipulation: catching a single molecule of DNA, RNA or a single protein and applying mechanical constrains using micron-size magnetic beads and magnets turn out to be easy. Various factors have made this possible: the fact that manufacturers have been preparing these beads to catch various biological entities-the ease of use provided by magnets which apply a force or a torque at a distance thus inside a flow cell-some chance: since the forces so generated are in the right range to stretch a single molecule. This is a little less true for torque. Finally, one feature which also appears very important is the simplicity of their calibration using Brownian motion. Here we start by describing magnetic tweezers used routinely in our laboratory where we have tried to develop a device as simple as possible so that the experimentalist can really focus on the biological aspect of the biomolecules that he/she is interested in. We discuss the implications of the various components and their important features. Next, we summarize what is easy to achieve and what is less easy. Then we refer to contributions by other groups who have brought valuable insights to improve magnetic tweezers.

Dynamic 1D Search and Processive Nucleosome Translocations by RSC and ISW2 Chromatin Remodelers

Eukaryotic gene expression is linked to chromatin structure and nucleosome positioning by ATP-dependent chromatin remodelers that establish and maintain nucleosome-depleted regions (NDRs) near transcription start-sites. Conserved yeast RSC and ISW2 remodelers exert antagonistic effects on nucleosomes flanking NDRs, but the temporal dynamics of remodeler search, engagement and directional nucleosome mobilization for promoter accessibility are unknown. Using optical tweezers and 2-color single-particle imaging, we investigated the Brownian diffusion of RSC and ISW2 on free DNA and sparse nucleosome arrays. RSC and ISW2 rapidly scan DNA by one-dimensional hopping and sliding respectively, with dynamic collisions between remodelers followed by recoil or apparent co-diffusion. Static nucleosomes block remodeler diffusion resulting in remodeler recoil or sequestration. Remarkably, both RSC and ISW2 use ATP hydrolysis to translocate mono-nucleosomes processively at ~30 bp/sec on extended linear DNA under tension. Processivity and opposing push-pull directionalities of nucleosome translocation shown by RSC and ISW2 shape the distinctive landscape of promoter chromatin.

Correlated Single-Molecule Magnetic Tweezers and Fluorescence Measurements of DNA-Enzyme Interactions

Combining force spectroscopy and fluorescence microscopy provides a substantial improvement to the single-molecule toolbox by allowing simultaneous manipulation and orthogonal characterizations of the conformations, interactions, and activity of biomolecular complexes. Here, we describe a combined magnetic tweezers and total internal reflection fluorescence microscopy setup to carry out correlated single-molecule fluorescence spectroscopy and force/twisting experiments. We apply the setup to reveal the DNA interactions of the CRISPR-Cas surveillance complex Cascade. Single-molecule fluorescence of a labeled Cascade allows to follow the DNA association and dissociation of the protein. Simultaneously, the magnetic tweezers probe the DNA unwinding during R-loop formation by the bound Cascade complexes. Furthermore, the setup supports observation of 1D diffusion of protein complexes on stretched DNA molecules. This technique can be applied to study a vast range of protein-DNA interactions.

Trends in Single-Molecule Total Internal Reflection Fluorescence Imaging and Their Biological Applications with Lab-on-a-Chip Technology

Single-molecule imaging technologies, especially those based on fluorescence, have been developed to probe both the equilibrium and dynamic properties of biomolecules at the single-molecular and quantitative levels. In this review, we provide an overview of the state-of-the-art advancements in single-molecule fluorescence imaging techniques. We systematically explore the advanced implementations of in vitro single-molecule imaging techniques using total internal reflection fluorescence (TIRF) microscopy, which is widely accessible. This includes discussions on sample preparation, passivation techniques, data collection and analysis, and biological applications. Furthermore, we delve into the compatibility of microfluidic technology for single-molecule fluorescence imaging, highlighting its potential benefits and challenges. Finally, we summarize the current challenges and prospects of fluorescence-based single-molecule imaging techniques, paving the way for further advancements in this rapidly evolving field.

Single-molecule study reveals Hmo1, not Hho1, promotes chromatin assembly in budding yeast

Linker histone H1 plays a crucial role in various biological processes, including nucleosome stabilization, high-order chromatin structure organization, gene expression, and epigenetic regulation in eukaryotic cells. Unlike higher eukaryotes, little about the linker histone in Saccharomyces cerevisiae is known. Hho1 and Hmo1 are two long-standing controversial histone H1 candidates in budding yeast. In this study, we directly observed at the single-molecule level that Hmo1, but not Hho1, is involved in chromatin assembly in the yeast nucleoplasmic extracts (YNPE), which can replicate the physiological condition of the yeast nucleus. The presence of Hmo1 facilitates the assembly of nucleosomes on DNA in YNPE, as revealed by single-molecule force spectroscopy. Further single-molecule analysis showed that the lysine-rich C-terminal domain (CTD) of Hmo1 is essential for the function of chromatin compaction, while the second globular domain at the C-terminus of Hho1 impairs its ability. In addition, Hmo1, but not Hho1, forms condensates with double-stranded DNA via reversible phase separation. The phosphorylation fluctuation of Hmo1 coincides with metazoan H1 during the cell cycle. Our data suggest that Hmo1, but not Hho1, possesses some functionality similar to that of linker histone in Saccharomyces cerevisiae, even though some properties of Hmo1 differ from those of a canonical linker histone H1. Our study provides clues for the linker histone H1 in budding yeast and provides insights into the evolution and diversity of histone H1 across eukaryotes. IMPORTANCE There has been a long-standing debate regarding the identity of linker histone H1 in budding yeast. To address this issue, we utilized YNPE, which accurately replicate the physiological conditions in yeast nuclei, in combination with total internal reflection fluorescence microscopy and magnetic tweezers. Our findings demonstrated that Hmo1, rather than Hho1, is responsible for chromatin assembly in budding yeast. Additionally, we found that Hmo1 shares certain characteristics with histone H1, including phase separation and phosphorylation fluctuations throughout the cell cycle. Furthermore, we discovered that the lysine-rich domain of Hho1 is buried by its second globular domain at the C-terminus, resulting in the loss of function that is similar to histone H1. Our study provides compelling evidence to suggest that Hmo1 shares linker histone H1 function in budding yeast and contributes to our understanding of the evolution of linker histone H1 across eukaryotes.

ATP binding facilitates target search of SWR1 chromatin remodeler by promoting one-dimensional diffusion on DNA

One-dimensional (1D) target search is a well-characterized phenomenon for many DNA-binding proteins but is poorly understood for chromatin remodelers. Herein, we characterize the 1D scanning properties of SWR1, a conserved yeast chromatin remodeler that performs histone exchange on +1 nucleosomes adjacent to a nucleosome-depleted region (NDR) at gene promoters. We demonstrate that SWR1 has a kinetic binding preference for DNA of NDR length as opposed to gene-body linker length DNA. Using single and dual color single-particle tracking on DNA stretched with optical tweezers, we directly observe SWR1 diffusion on DNA. We found that various factors impact SWR1 scanning, including ATP which promotes diffusion through nucleotide binding rather than ATP hydrolysis. A DNA-binding subunit, Swc2, plays an important role in the overall diffusive behavior of the complex, as the subunit in isolation retains similar, although faster, scanning properties as the whole remodeler. ATP-bound SWR1 slides until it encounters a protein roadblock, of which we tested dCas9 and nucleosomes. The median diffusion coefficient, 0.024 μm2/s, in the regime of helical sliding, would mediate rapid encounter of NDR-flanking nucleosomes at length scales found in cellular chromatin.

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.

How p53 Molecules Solve the Target DNA Search Problem: A Review

Interactions between DNA and DNA-binding proteins play an important role in many essential cellular processes. A key function of the DNA-binding protein p53 is to search for and bind to target sites incorporated in genomic DNA, which triggers transcriptional regulation. How do p53 molecules achieve "rapid" and "accurate" target search in living cells? The search dynamics of p53 were expected to include 3D diffusion in solution, 1D diffusion along DNA, and intersegmental transfer between two different DNA strands. Single-molecule fluorescence microscopy enabled the tracking of p53 molecules on DNA and the characterization of these dynamics quantitatively. Recent intensive single-molecule studies of p53 succeeded in revealing each of these search dynamics. Here, we review these studies and discuss the target search mechanisms of p53.

Single-molecule visualization reveals the damage search mechanism for the human NER protein XPC-RAD23B

DNA repair is critical for maintaining genomic integrity. Finding DNA lesions initiates the entire repair process. In human nucleotide excision repair (NER), XPC-RAD23B recognizes DNA lesions and recruits downstream factors. Although previous studies revealed the molecular features of damage identification by the yeast orthologs Rad4-Rad23, the dynamic mechanisms by which human XPC-RAD23B recognizes DNA defects have remained elusive. Here, we directly visualized the motion of XPC-RAD23B on undamaged and lesion-containing DNA using high-throughput single-molecule imaging. We observed three types of one-dimensional motion of XPC-RAD23B along DNA: diffusive, immobile and constrained. We found that consecutive AT-tracks led to increase in proteins with constrained motion. The diffusion coefficient dramatically increased according to ionic strength, suggesting that XPC-RAD23B diffuses along DNA via hopping, allowing XPC-RAD23B to bypass protein obstacles during the search for DNA damage. We also examined how XPC-RAD23B identifies cyclobutane pyrimidine dimers (CPDs) during diffusion. XPC-RAD23B makes futile attempts to bind to CPDs, consistent with low CPD recognition efficiency. Moreover, XPC-RAD23B binds CPDs in biphasic states, stable for lesion recognition and transient for lesion interrogation. Taken together, our results provide new insight into how XPC-RAD23B searches for DNA lesions in billions of base pairs in human genome.

Intrinsically disordered regions regulate both catalytic and non-catalytic activities of the MutLα mismatch repair complex

Intrinsically disordered regions (IDRs) are present in at least 30% of the eukaryotic proteome and are enriched in chromatin-associated proteins. Using a combination of genetics, biochemistry and single-molecule biophysics, we characterize how IDRs regulate the functions of the yeast MutLα (Mlh1-Pms1) mismatch repair (MMR) complex. Shortening or scrambling the IDRs in both subunits ablates MMR in vivo. Mlh1-Pms1 complexes with shorter IDRs that disrupt MMR retain wild-type DNA binding affinity but are impaired for diffusion on both naked and nucleosome-coated DNA. Moreover, the IDRs also regulate the adenosine triphosphate hydrolysis and nuclease activities that are encoded in the structured N- and C-terminal domains of the complex. This combination of phenotypes underlies the catastrophic MMR defect seen with the mutant MutLα in vivo. More broadly, this work highlights an unanticipated multi-functional role for IDRs in regulating both facilitated diffusion on chromatin and nucleolytic processing of a DNA substrate.

High Free-Energy Barrier of 1D Diffusion Along DNA by Architectural DNA-Binding Proteins

Architectural DNA-binding proteins function to regulate diverse DNA reactions and have the defining property of significantly changing DNA conformation. Although the 1D movement along DNA by other types of DNA-binding proteins has been visualized, the mobility of architectural DNA-binding proteins on DNA remains unknown. Here, we applied single-molecule fluorescence imaging on arrays of extended DNA molecules to probe the binding dynamics of three structurally distinct architectural DNA-binding proteins: Nhp6A, HU, and Fis. Each of these proteins was observed to move along DNA, and the salt concentration independence of the 1D diffusion implies sliding with continuous contact to DNA. Nhp6A and HU exhibit a single sliding mode, whereas Fis exhibits two sliding modes. Based on comparison of the diffusion coefficients and sizes of many DNA binding proteins, the architectural proteins are categorized into a new group distinguished by an unusually high free-energy barrier for 1D diffusion. The higher free-energy barrier for 1D diffusion by architectural proteins can be attributed to the large DNA conformational changes that accompany binding and impede rotation-coupled movement along the DNA grooves.

The more the merrier: high-throughput single-molecule techniques

The single-molecule approach seeks to understand molecular mechanisms by observing biomolecular processes at the level of individual molecules. These methods have led to a developing understanding that for many processes, a diversity of behaviours will be observed, representing a multitude of pathways. This realisation necessitates that an adequate number of observations are recorded to fully characterise this diversity. The requirement for large numbers of observations to adequately sample distributions, subpopulations, and rare events presents a significant challenge for single-molecule techniques, which by their nature do not typically provide very high throughput. This review will discuss many developing techniques which address this issue by combining nanolithographic approaches, such as zero-mode waveguides and DNA curtains, with single-molecule fluorescence microscopy, and by drastically increasing throughput of force-based approaches such as magnetic tweezers and laminar-flow techniques. These methods not only allow the collection of large volumes of single-molecule data in single experiments, but have also made improvements to ease-of-use, accessibility, and automation of data analysis.

Hide and seek: How do DNA glycosylases locate oxidatively damaged DNA bases amidst a sea of undamaged bases?

The first step of the base excision repair (BER) pathway responsible for removing oxidative DNA damage utilizes DNA glycosylases to find and remove the damaged DNA base. How glycosylases find the damaged base amidst a sea of undamaged bases has long been a question in the BER field. Single molecule total internal reflection fluorescence microscopy (SM TIRFM) experiments have allowed for an exciting look into this search mechanism and have found that DNA glycosylases scan along the DNA backbone in a bidirectional and random fashion. By comparing the search behavior of bacterial glycosylases from different structural families and with varying substrate specificities, it was found that glycosylases search for damage by periodically inserting a wedge residue into the DNA stack as they redundantly search tracks of DNA that are 450-600bp in length. These studies open up a wealth of possibilities for further study in real time of the interactions of DNA glycosylases and other BER enzymes with various DNA substrates.

Single-Molecule Methods for Nucleotide Excision Repair: Building a System to Watch Repair in Real Time

Single-molecule approaches to solving biophysical problems are powerful tools that allow static and dynamic real-time observations of specific molecular interactions of interest in the absence of ensemble-averaging effects. Here, we provide detailed protocols for building an experimental system that employs atomic force microscopy and a single-molecule DNA tightrope assay based on oblique angle illumination fluorescence microscopy. Together with approaches for engineering site-specific lesions into DNA substrates, these complementary biophysical techniques are well suited for investigating protein-DNA interactions that involve target-specific DNA-binding proteins, such as those engaged in a variety of DNA repair pathways. In this chapter, we demonstrate the utility of the platform by applying these techniques in the studies of proteins participating in nucleotide excision repair.

Using single-molecule approach to visualize the nucleosome assembly in yeast nucleoplasmic extracts

In eukaryotic cells, the smallest subunit of chromatin is the nucleosome, which consists of a segment of DNA wound on histone protein cores. Despite many years of effort, the process of nucleosome assembly and disassembly is still not very clear. Here, we present a convenient method to investigate the process of nucleosome assembly at the single molecule level. We invented a novel system derived from the yeast nucleoplasmic extracts (YNPE), and demonstrated that the YNPE supports the nucleosome assembly under physiological condition. By combining the total internal reflection fluorescence microscopy with microfluidic flow-cell technique, the dynamic process of nucleosome assembly in YNPE was visualized at single-molecule level. Our system provides a novel in vitro single-molecule tool to investigate the dynamics of nucleosome assembly under physiological conditions.

Chemical Biology Approaches to Genome Editing: Understanding, Controlling, and Delivering Programmable Nucleases

Programmable DNA nucleases have provided scientists with the unprecedented ability to probe, regulate, and manipulate the human genome. Zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and the clustered regularly interspaced short palindromic repeat-Cas9 system (CRISPR-Cas9) represent a powerful array of tools that can bind to and cleave a specified DNA sequence. In their canonical forms, these nucleases induce double-strand breaks at a DNA locus of interest that can trigger cellular DNA repair processes that disrupt or replace genes. The fusion of these programmable nucleases with a variety of other protein domains has led to a rapidly growing suite of tools for activating, repressing, visualizing, and modifying loci of interest. Maximizing the usefulness and therapeutic relevance of these tools, however, requires precisely controlling their activity and specificity to minimize potentially toxic side effects arising from off-target activities. This need has motivated the application of chemical biology principles and methods to genome-editing proteins, including the engineering of variants of these proteins with improved or altered specificities, and the development of genetic, chemical, optical, and protein delivery methods that control the activity of these agents in cells. Advancing the capabilities, safety, effectiveness, and therapeutic relevance of genome-engineering proteins will continue to rely on chemical biology strategies that manipulate their activity, specificity, and localization.

Dynamic Chromatin Regulation from a Single Molecule Perspective

Chromatin regulatory processes, like all biological reactions, are dynamic and stochastic in nature but can give rise to stable and inheritable changes in gene expression patterns. A molecular understanding of those processes is key for fundamental biological insight into gene regulation, epigenetic inheritance, lineage determination, and therapeutic intervention in the case of disease. In recent years, great progress has been made in identifying important molecular players involved in key chromatin regulatory pathways. Conversely, we are only beginning to understand the dynamic interplay between protein effectors, transcription factors, and the chromatin substrate itself. Single-molecule approaches employing both highly defined chromatin substrates in vitro, as well as direct observation of complex regulatory processes in vivo, open new avenues for a molecular view of chromatin regulation. This review highlights recent applications of single-molecule methods and related techniques to investigate fundamental chromatin regulatory processes.

Analysis of Combinatorial Epigenomic States

Hundreds of distinct chemical modifications to DNA and histone amino acids have been described. Regulation exerted by these so-called epigenetic marks is vital to normal development, stability of cell identity through mitosis, and nongenetic transmission of traits between generations through meiosis. Loss of this regulation contributes to many diseases. Evidence indicates epigenetic marks function in combinations, whereby a given modification has distinct effects on local genome control, depending on which additional modifications are locally present. This review summarizes emerging methods for assessing combinatorial epigenomic states, as well as challenges and opportunities for their refinement.

Visualization of large elongated DNA molecules

Long and linear DNA molecules are the mainstream single-molecule analytes for a variety of biochemical analysis within microfluidic devices, including functionalized surfaces and nanostructures. However, for biochemical analysis, large DNA molecules have to be unraveled, elongated, and visualized to obtain biochemical and genomic information. To date, elongated DNA molecules have been exploited in the development of a number of genome analysis systems as well as for the study of polymer physics due to the advantage of direct visualization of single DNA molecule. Moreover, each single DNA molecule provides individual information, which makes it useful for stochastic event analysis. Therefore, numerous studies of enzymatic random motions have been performed on a large elongated DNA molecule. In this review, we introduce mechanisms to elongate DNA molecules using microfluidics and nanostructures in the beginning. Secondly, we discuss how elongated DNA molecules have been utilized to obtain biochemical and genomic information by direct visualization of DNA molecules. Finally, we reviewed the approaches used to study the interaction of proteins and large DNA molecules. Although DNA-protein interactions have been investigated for many decades, it is noticeable that there have been significant achievements for the last five years. Therefore, we focus mainly on recent developments for monitoring enzymatic activity on large elongated DNA molecules.

Using supported bilayers to study the spatiotemporal organization of membrane-bound proteins

Cell division in prokaryotes and eukaryotes is commonly initiated by the well-controlled binding of proteins to the cytoplasmic side of the cell membrane. However, a precise characterization of the spatiotemporal dynamics of membrane-bound proteins is often difficult to achieve in vivo. Here, we present protocols for the use of supported lipid bilayers to rebuild the cytokinetic machineries of cells with greatly different dimensions: the bacterium Escherichia coli and eggs of the vertebrate Xenopus laevis. Combined with total internal reflection fluorescence microscopy, these experimental setups allow for precise quantitative analyses of membrane-bound proteins. The protocols described to obtain glass-supported membranes from bacterial and vertebrate lipids can be used as starting points for other reconstitution experiments. We believe that similar biochemical assays will be instrumental to study the biochemistry and biophysics underlying a variety of complex cellular tasks, such as signaling, vesicle trafficking, and cell motility.

Single molecule fluorescence methodologies for investigating transcription factor binding kinetics to nucleosomes and DNA

Site specific DNA binding complexes must bind their DNA target sites and then reside there for a sufficient amount of time for proper regulation of DNA processing including transcription, replication and DNA repair. In eukaryotes, the occupancy of DNA binding complexes at their target sites is regulated by chromatin structure and dynamics. Methodologies that probe both the binding and dissociation kinetics of DNA binding proteins with naked and nucleosomal DNA are essential for understanding the mechanisms by which these complexes function. Here, we describe single-molecule fluorescence methodologies for quantifying the binding and dissociation kinetics of transcription factors at a target site within DNA, nucleosomes and nucleosome arrays. This approach allowed for the unexpected observation that nucleosomes impact not only binding but also dissociation kinetics of transcription factors and is well-suited for the investigation of numerous DNA processing complexes that directly interact with DNA organized into chromatin.

Single molecule and single cell epigenomics

Dynamically regulated changes in chromatin states are vital for normal development and can produce disease when they go awry. Accordingly, much effort has been devoted to characterizing these states under normal and pathological conditions. Chromatin immunoprecipitation followed by sequencing (ChIP-seq) is the most widely used method to characterize where in the genome transcription factors, modified histones, modified nucleotides and chromatin binding proteins are found; bisulfite sequencing (BS-seq) and its variants are commonly used to characterize the locations of DNA modifications. Though very powerful, these methods are not without limitations. Notably, they are best at characterizing one chromatin feature at a time, yet chromatin features arise and function in combination. Investigators commonly superimpose separate ChIP-seq or BS-seq datasets, and then infer where chromatin features are found together. While these inferences might be correct, they can be misleading when the chromatin source has distinct cell types, or when a given cell type exhibits any cell to cell variation in chromatin state. These ambiguities can be eliminated by robust methods that directly characterize the existence and genomic locations of combinations of chromatin features in very small inputs of cells or ideally, single cells. Here we review single molecule epigenomic methods under development to overcome these limitations, the technical challenges associated with single molecule methods and their potential application to single cells.

Rapid prototyping of multichannel microfluidic devices for single-molecule DNA curtain imaging

Single-molecule imaging and manipulation of biochemical reactions continues to reveal numerous biological insights. To facilitate these studies, we have developed and implemented a high-throughput approach to organize and image hundreds of individual DNA molecules at aligned diffusion barriers. Nonetheless, obtaining statistically relevant data sets under a variety of reaction conditions remains challenging. Here, we present a method for integrating high-throughput single-molecule "DNA curtain" imaging with poly(dimethylsiloxane) (PDMS)-based microfluidics. Our benchtop fabrication method can be accomplished in minutes with common tools found in all molecular biology laboratories. We demonstrate the utility of this approach by simultaneous imaging of two independent biochemical reaction conditions in a laminar flow device. In addition, five different reaction conditions can be observed concurrently in a passive linear gradient generator. Combining rapid microfluidic fabrication with high-throughput DNA curtains greatly expands our capability to interrogate complex biological reactions.

Insights into the glycosylase search for damage from single-molecule fluorescence microscopy

The first step of base excision repair utilizes glycosylase enzymes to find damage within a genome. A persistent question in the field of DNA repair is how glycosylases interact with DNA to specifically find and excise target damaged bases with high efficiency and specificity. Ensemble studies have indicated that glycosylase enzymes rely upon both sliding and distributive modes of search, but ensemble methods are limited in their ability to directly observe these modes. Here we review insights into glycosylase scanning behavior gathered through single-molecule fluorescence studies of enzyme interactions with DNA and provide a context for these results in relation to ensemble experiments.

DNA interrogation by the CRISPR RNA-guided endonuclease Cas9

The CRISPR-associated enzyme Cas9 is an RNA-guided endonuclease that uses RNA:DNA base-pairing to target foreign DNA in bacteria. Cas9:guide RNA complexes are also effective genome engineering agents in animals and plants. Here we use single-molecule and bulk biochemical experiments to determine how Cas9:RNA interrogates DNA to find specific cleavage sites. We show that both binding and cleavage of DNA by Cas9:RNA require recognition of a short trinucleotide protospacer adjacent motif (PAM). Non-target DNA binding affinity scales with PAM density, and sequences fully complementary to the guide RNA but lacking a nearby PAM are ignored by Cas9:RNA. DNA strand separation and RNA:DNA heteroduplex formation initiate at the PAM and proceed directionally towards the distal end of the target sequence. Furthermore, PAM interactions trigger Cas9 catalytic activity. These results reveal how Cas9 employs PAM recognition to quickly identify potential target sites while scanning large DNA molecules, and to regulate dsDNA scission.

Toward single-molecule optical mapping of the epigenome

The past decade has seen an explosive growth in the utilization of single-molecule techniques for the study of complex systems. The ability to resolve phenomena otherwise masked by ensemble averaging has made these approaches especially attractive for the study of biological systems, where stochastic events lead to inherent inhomogeneity at the population level. The complex composition of the genome has made it an ideal system to study at the single-molecule level, and methods aimed at resolving genetic information from long, individual, genomic DNA molecules have been in use for the last 30 years. These methods, and particularly optical-based mapping of DNA, have been instrumental in highlighting genomic variation and contributed significantly to the assembly of many genomes including the human genome. Nanotechnology and nanoscopy have been a strong driving force for advancing genomic mapping approaches, allowing both better manipulation of DNA on the nanoscale and enhanced optical resolving power for analysis of genomic information. During the past few years, these developments have been adopted also for epigenetic studies. The common principle for these studies is the use of advanced optical microscopy for the detection of fluorescently labeled epigenetic marks on long, extended DNA molecules. Here we will discuss recent single-molecule studies for the mapping of chromatin composition and epigenetic DNA modifications, such as DNA methylation.

Single-molecule and single-particle imaging of molecular motors in vitro and in vivo

Motor proteins are multi-potent molecular machines, whose localisation, function and regulation are achieved through tightly controlled processes involving conformational changes and interactions with their tracks, cargos and binding partners. Understanding how these complex machines work requires dissection of these processes both in space and time. Complementing the traditional ensemble measurements, single-molecule assays enable the detection of rare or short-lived intermediates and molecular heterogeneities, and the measurements of subpopulation dynamics. This chapter is focusing on the fluorescence imaging of single motors and their cargo. It discusses what is required in order to achieve single-molecule imaging with high temporal and spatial resolution and how these requirements are met both in vitro and in vivo. It also presents a general overview and applied examples of the major single-molecule imaging techniques and experimental assays which have been used to study motor proteins.

Micro- and nanoscale devices for the investigation of epigenetics and chromatin dynamics

Deoxyribonucleic acid (DNA) is the blueprint on which life is based and transmitted, but the way in which chromatin - a dynamic complex of nucleic acids and proteins - is packaged and behaves in the cellular nucleus has only begun to be investigated. Epigenetic modifications sit 'on top of' the genome and affect how DNA is compacted into chromatin and transcribed into ribonucleic acid (RNA). The packaging and modifications around the genome have been shown to exert significant influence on cellular behaviour and, in turn, human development and disease. However, conventional techniques for studying epigenetic or conformational modifications of chromosomes have inherent limitations and, therefore, new methods based on micro- and nanoscale devices have been sought. Here, we review the development of these devices and explore their use in the study of DNA modifications, chromatin modifications and higher-order chromatin structures.

Lab-on-a-chip technologies for single-molecule studies

Recent developments on various lab-on-a-chip techniques allow miniaturized and integrated devices to perform on-chip single-molecule studies. Fluidic-based platforms that utilize unique microscale fluidic behavior are capable of conducting single-molecule experiments with high sensitivities and throughputs, while biomolecular systems can be studied on-chip using techniques such as DNA curtains, magnetic tweezers, and solid-state nanopores. The advances of these on-chip single-molecule techniques lead to next-generation lab-on-a-chip devices, such as DNA transistors, and single-molecule real-time (SMRT) technology for rapid and low-cost whole genome DNA sequencing. In this Focus article, we will discuss some recent successes in the development of lab-on-a-chip techniques for single-molecule studies and expound our thoughts on the near future of on-chip single-molecule studies.

Archaeal nucleosome positioning in vivo and in vitro is directed by primary sequence motifs

Background

Histone wrapping of DNA into nucleosomes almost certainly evolved in the Archaea, and predates Eukaryotes. In Eukaryotes, nucleosome positioning plays a central role in regulating gene expression and is directed by primary sequence motifs that together form a nucleosome positioning code. The experiments reported were undertaken to determine if archaeal histone assembly conforms to the nucleosome positioning code.

Results

Eukaryotic nucleosome positioning is favored and directed by phased helical repeats of AA/TT/AT/TA and CC/GG/CG/GC dinucleotides, and disfavored by longer AT-rich oligonucleotides. Deep sequencing of genomic DNA protected from micrococcal nuclease digestion by assembly into archaeal nucleosomes has established that archaeal nucleosome assembly is also directed and positioned by these sequence motifs, both in vivo in Methanothermobacter thermautotrophicus and Thermococcus kodakarensis and in vitro in reaction mixtures containing only one purified archaeal histone and genomic DNA. Archaeal nucleosomes assembled at the same locations in vivo and in vitro, with much reduced assembly immediately upstream of open reading frames and throughout the ribosomal rDNA operons. Providing further support for a common positioning code, archaeal histones assembled into nucleosomes on eukaryotic DNA and eukaryotic histones into nucleosomes on archaeal DNA at the same locations. T. kodakarensis has two histones, designated HTkA and HTkB, and strains with either but not both histones deleted grow normally but do exhibit transcriptome differences. Comparisons of the archaeal nucleosome profiles in the intergenic regions immediately upstream of genes that exhibited increased or decreased transcription in the absence of HTkA or HTkB revealed substantial differences but no consistent pattern of changes that would correlate directly with archaeal nucleosome positioning inhibiting or stimulating transcription.

Conclusions

The results obtained establish that an archaeal histone and a genome sequence together are sufficient to determine where archaeal nucleosomes preferentially assemble and where they avoid assembly. We confirm that the same nucleosome positioning code operates in Archaea as in Eukaryotes and presumably therefore evolved with the histone-fold mechanism of DNA binding and compaction early in the archaeal lineage, before the divergence of Eukaryotes.

Towards physiological complexity with in vitro single-molecule biophysics

Single-molecule biology has matured in recent years, driven to greater sophistication by the development of increasingly advanced experimental techniques. A progressive appreciation for its unique strengths is attracting research that spans an exceptionally broad swath of physiological phenomena--from the function of nucleosomes to protein diffusion in the cell membrane. Newfound enthusiasm notwithstanding, the single-molecule approach is limited to an intrinsically defined set of biological questions; such limitation applies to all experimental approaches, and an explicit statement of the boundaries delineating each set offers a guide to most fruitfully orienting in vitro single-molecule research in the future. Here, we briefly describe a simple conceptual framework to categorize how submolecular, molecular and intracellular processes are studied. We highlight the domain of single-molecule biology in this scheme, with an emphasis on its ability to probe various forms of heterogeneity inherent to populations of discrete biological macromolecules. We then give a general overview of our high-throughput DNA curtain methodology for studying protein-nucleic acid interactions, and by contextualizing it within this framework, we explore what might be the most enticing avenues of future research. We anticipate that a focus on single-molecule biology's unique strengths will suggest a new generation of experiments with greater complexity and more immediately translatable physiological relevance.

The conserved PHD1-PHD2 domain of ZFP-1/AF10 is a discrete functional module essential for viability in Caenorhabditis elegans

Plant homeodomain (PHD)-type zinc fingers play an important role in recognizing chromatin modifications and recruiting regulatory proteins to specific genes. A specific module containing a conventional PHD finger followed by an extended PHD finger exists in the mammalian AF10 protein, among a few others. AF10 has mostly been studied in the context of the leukemic MLL-AF10 fusion protein, which lacks the N-terminal PHD fingers of AF10. Although this domain of AF10 is the most conserved region of the protein, its biological significance has not been elucidated. In this study, we used genetic and biochemical approaches to examine the PHD1-PHD2 region of the Caenorhabditis elegans ortholog of AF10, zinc finger protein 1 (ZFP-1). We demonstrate that the PHD1-PHD2 region is essential for viability and that the first PHD finger contributes to the preferred binding of PHD1-PHD2 to lysine 4-methylated histone H3 tails. Moreover, we show that ZFP-1 localization peaks overlap with H3K4 methylation-enriched promoters of actively expressed genes genomewide and that H3K4 methylation is important for ZFP-1 localization to promoters in the embryo. We predict that the essential biological role of the PHD1-PHD2 module of ZFP-1/AF10 is connected to the regulation of actively expressed genes during early development.

Single-stranded DNA curtains for real-time single-molecule visualization of protein-nucleic acid interactions

Single-molecule imaging of biological macromolecules has dramatically impacted our understanding of many types of biochemical reactions. To facilitate these studies, we have established new strategies for anchoring and organizing DNA molecules on the surfaces of microfluidic sample chambers that are otherwise coated with fluid lipid bilayers. This previous work was reliant upon the use of double-stranded DNA, precluding access to information on biological processes involving single-stranded nucleic acid substrates. Here, we present procedures for aligning and visualizing single-stranded DNA molecules along the leading edges of nanofabricated barriers to lipid diffusion, in both "single-tethered" and "double-tethered" experimental formats. This new single-molecule approach provides long-awaited access to critical biological reactions involving single-stranded DNA binding proteins.

Measuring intermolecular rupture forces with a combined TIRF-optical trap microscope and DNA curtains

We report a new approach to probing DNA-protein interactions by combining optical tweezers with a high-throughput DNA curtains technique. Here we determine the forces required to remove the individual lipid-anchored DNA molecules from the bilayer. We demonstrate that DNA anchored to the bilayer through a single biotin-streptavidin linkage withstands ∼20pN before being pulled free from the bilayer, whereas molecules anchored to the bilayer through multiple attachment points can withstand ⩾65pN; access to this higher force regime is sufficient to probe the responses of protein-DNA interactions to force changes. As a proof-of-principle, we concurrently visualized DNA-bound fluorescently-tagged RNA polymerase while simultaneously stretching the DNA molecules. This work presents a step towards a powerful experimental platform that will enable concurrent visualization of DNA curtains while applying defined forces through optical tweezers.

Assembly of DNA curtains using hydrogen silsesquioxane as a barrier to lipid diffusion

We have established a single-molecule imaging experimental platform called "DNA curtains" in which DNA molecules tethered to a lipid bilayer are organized into patterns at nanofabricated metallic barriers on the surface of a microfluidic sample chamber. This technology has wide applications for real-time single-molecule imaging of protein-nucleic acid interactions. Here, we demonstrate that DNA curtains can also be made from hydrogen silsesquioxane (HSQ). HSQ offers important advantages over metallic barriers because it can be lithographically patterned directly onto fused silica slides without any requirement for further processing steps, thereby offering the potential for rapid prototype development and/or scale up for manufacturing.

DNA structure, nucleosome placement and chromatin remodelling: a perspective

A major question in chromatin biology is to what extent the sequence of DNA directly determines the genetic and chromatin organization of a eukaryotic genome? We consider two aspects to this question: the DNA sequence-specified positioning of nucleosomes and the determination of NDRs (nucleosome-depleted regions) or barriers. We argue that, in budding yeast, while DNA sequence-specified nucleosome positioning may contribute to positions flanking the regions lacking nucleosomes, DNA thermodynamic stability is a major component determinant of the genetic organization of this organism.

Detection of nucleosomal substructures using solid-state nanopores

Histone proteins assemble onto DNA into nucleosomes that control the structure and function of eukaryotic chromatin. More specifically, the structural integrity of nucleosomes regulates gene expression rates and serves as an important early marker for cell apoptosis. Nucleosomal (sub)structures are however hard to detect and characterize. Here, we show that solid-state nanopores are well suited for fast and label-free detection of nucleosomes and its histone subcomplexes. (Nucleo-)protein complexes are individually driven through the nanopore by an applied electric field, which results in characteristic conductance blockades that provide quantitative information on the molecular size of the translocating complex. We observe a systematic dependence of the conductance blockade and translocation time on the molecular weight of the nucleosomal substructures. This allows discriminating and characterizing these protein and DNA-protein complexes at the single-complex level. Finally, we demonstrate the ability to distinguish nucleosomes and dinucleosomes as a first step toward using the nanopore platform to study chromatin arrays.

Recent advances in single molecule studies of nucleosomes

As the fundamental packing units of DNA in eukaryotes, nucleosomes play a central role in governing DNA accessibility in a variety of cellular processes. Our understanding of the mechanisms underlying this complex regulation has been aided by unique structural and dynamic perspectives offered by single molecule techniques. Recent years have witnessed remarkable advances achieved using these techniques, including the generation of a detailed histone-DNA energy landscape, elucidation of nucleosome disassembly processes, and real-time monitoring of molecular motors interacting with nucleosomes. These and other highlights of single molecule nucleosome studies will be discussed in this review.

Nanofabrication for the analysis and manipulation of membranes

Recent advancements and applications of nanofabrication have enabled the characterization and control of biological membranes at submicron scales. This review focuses on the application of nanofabrication towards the nanoscale observing, patterning, sorting, and concentrating membrane components. Membranes on living cells are a necessary component of many fundamental cellular processes that naturally incorporate nanoscale rearrangement of the membrane lipids and proteins. Nanofabrication has advanced these understandings, for example, by providing 30 nm resolution of membrane proteins with metal-enhanced fluorescence at the tip of a scanning probe on fixed cells. Naturally diffusing single molecules at high concentrations on live cells have been observed at 60 nm resolution by confining the fluorescence excitation light through nanoscale metallic apertures. The lateral reorganization on the plasma membrane during membrane-mediated signaling processes has been examined in response to nanoscale variations in the patterning and mobility of the signal-triggering molecules. Further, membrane components have been separated, concentrated, and extracted through on-chip electrophoretic and microfluidic methods. Nanofabrication provides numerous methods for examining and manipulating membranes for both greater understandings of membrane processes as well as for the application of membranes to other biophysical methods.

Quantum dots find their stride in single molecule tracking

Thirteen years after the demonstration of quantum dots as biological imaging agents, and nine years after the initial commercial introduction of bioconjugated quantum dots, the brightness and photostability of the quantum dots has enabled a range of investigations using single molecule tracking. These materials are being routinely utilized by a number of groups to track the dynamics of single molecules in reconstituted biophysical systems and on living cells, and are especially powerful for investigations of single molecules over long timescales with short exposure times and high pointing accuracy. New approaches are emerging where the quantum dots are used as 'hard-sphere' probes for intracellular compartments. Innovations in quantum dot surface modification are poised to substantially expand the utility of these materials.

Nonhistone Scm3 binds to AT-rich DNA to organize atypical centromeric nucleosome of budding yeast

The molecular architecture of centromere-specific nucleosomes containing histone variant CenH3 is controversial. We have biochemically reconstituted two distinct populations of nucleosomes containing Saccharomyces cerevisiae CenH3 (Cse4). Reconstitution of octameric nucleosomes containing histones Cse4/H4/H2A/H2B is robust on noncentromere DNA, but inefficient on AT-rich centromere DNA. However, nonhistone Scm3, which is required for Cse4 deposition in vivo, facilitates in vitro reconstitution of Cse4/H4/Scm3 complexes on AT-rich centromere sequences. Scm3 has a nonspecific DNA binding domain that shows preference for AT-rich DNA and a histone chaperone domain that promotes specific loading of Cse4/H4. In live cells, Scm3-GFP is enriched at centromeres in all cell cycle phases. Chromatin immunoprecipitation confirms that Scm3 occupies centromere DNA throughout the cell cycle, even when Cse4 and H4 are temporarily dislodged in S phase. These findings suggest a model in which centromere-bound Scm3 aids recruitment of Cse4/H4 to assemble and maintain an H2A/H2B-deficient centromeric nucleosome.

Structure and Scm3-mediated assembly of budding yeast centromeric nucleosomes

Much controversy exists regarding the structural organization of the yeast centromeric nucleosome and the role of the nonhistone protein, Scm3, in its assembly and architecture. Here we show that the substitution of H3 with its centromeric variant Cse4 results in octameric nucleosomes that organize DNA in a left-handed superhelix. We demonstrate by single-molecule approaches, micrococcal nuclease digestion and small-angle X-ray scattering that Cse4-nucleosomes exhibit an open conformation with weakly bound terminal DNA segments. The Cse4-octamer does not preferentially form nucleosomes on its cognate centromeric DNA. We show that Scm3 functions as a Cse4-specific nucleosome assembly factor, and that the resulting octameric nucleosomes do not contain Scm3 as a stably bound component. Taken together, our data provide insights into the assembly and structural features of the budding yeast centromeric nucleosome.