At the core of molecular biology lies the intricate interplay between sequence, structure, and function. Single-molecule techniques provide in-depth dynamic insights into structure and function, but laborious assays impede functional screening of large sequence libraries. We introduce high-throughput Single-molecule Parallel Analysis for Rapid eXploration of Sequence space (SPARXS), integrating single-molecule fluorescence with next-generation sequencing. We applied SPARXS to study the sequence-dependent kinetics of the Holliday junction, a critical intermediate in homologous recombination. By examining the dynamics of millions of Holliday junctions, covering thousands of distinct sequences, we demonstrated the ability of SPARXS to uncover sequence patterns, evaluate sequence motifs, and construct thermodynamic models. SPARXS emerges as a versatile tool for untangling the mechanisms that underlie sequence-specific processes at the molecular scale.@en

Understanding the structure of biomolecules is vital for deciphering their roles in biological systems. Single-molecule techniques have emerged as alternatives to conventional ensemble structure analysis methods for uncovering new biology in molecular dynamics and interaction studies, yet only limited structural information could be obtained experimentally. Here, we address this challenge by introducing iMAX FRET, a one-pot method that allows ab initio 3D profiling of individual molecules using two-color FRET measurements. Through the stochastic exchange of fluorescent weak binders, iMAX FRET simultaneously assesses multiple distances on a biomolecule within a few minutes, which can then be used to reconstruct the coordinates of up to four points in each molecule, allowing structure-based inference. We demonstrate the 3D reconstruction of DNA nanostructures, protein quaternary structures, and conformational changes in proteins. With iMAX FRET, we provide a powerful approach to advance the understanding of biomolecular structure by expanding conventional FRET analysis to three dimensions.

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Proteins are the primary functional actors of the cell. While proteoform diversity is known to be highly biologically relevant, current protein analysis methods are of limited use for distinguishing proteoforms. Mass spectrometric methods, in particular, often provide only ambiguous information on post-translational modification sites, and sequences of co-existing modifications may not be resolved. Here we demonstrate fluorescence resonance energy transfer (FRET)-based single-molecule protein fingerprinting to map the location of individual amino acids and post-translational modifications within single full-length protein molecules. Our data show that both intrinsically disordered proteins and folded globular proteins can be fingerprinted with a subnanometer resolution, achieved by probing the amino acids one by one using single-molecule FRET via DNA exchange. This capability was demonstrated through the analysis of alpha-synuclein, an intrinsically disordered protein, by accurately quantifying isoforms in mixtures using a machine learning classifier, and by determining the locations of two O-GlcNAc moieties. Furthermore, we demonstrate fingerprinting of the globular proteins Bcl-2-like protein 1, procalcitonin and S100A9. We anticipate that our ability to perform proteoform identification with the ultimate sensitivity may unlock exciting new venues in proteomics research and biomarker-based diagnosis@en

The inherent properties of 2D materials—light mass, high out-of-plane flexibility, and large surface area—promise great potential for precise and accurate nanomechanical mass sensing, but their application is often hampered by surface contamination. Here we demonstrate a tri-layer graphene nanomechanical resonant mass sensor with sub-attogram resolution at room temperature, fabricated by a bottom-up process. We found that Joule-heating is effective in cleaning the graphene membrane surface, which results in a large improvement in the stability of the resonance frequency. We characterized the sensor by depositing Cr metal using a stencil mask and found a mass-resolution that is sufficient to weigh very small particles, like large proteins and protein complexes, with potential applications in the fields of nanobiology and medicine.

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Single-molecule fluorescence experiments allow monitoring of the structural change and dynamics of a single biomolecule in real time using dye molecules attached to the molecule. Often, the molecules are immobilized on the surface to observe a longer molecular dynamics, yet the finite photon budget available from an individual dye molecule before photobleaching sets the limit to the relatively poor signal-to-noise level. To increase the accuracy of these single-molecule experiments, it is necessary to study the cause of noise in the fluorescence signal from the single molecules. To find the origin of this noise, the lifetime of the fluorescent dye molecules labeled on surface-immobilized DNA was measured by using time-correlation single photon counting. The standard deviation of the fluorescence lifetimes obtained from repeated measurements of a single dye molecule with the total photon number N decreased as 1/N, thus following a shot noise of the Poisson statistics. On the other hand, an additional constant noise source, which is independent of the photon number, was observed from the lifetime uncertainties from many molecules and became more dominant after a certain photon number N. This trend was also followed in the uncertainties of the single-molecule FRET signals obtained from single and many molecules. This additional noise is considered to come from the inhomogeneous environment of each DNA immobilized on the surface.

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Single-cell profiling methods have had a profound impact on the understanding of cellular heterogeneity. While genomes and transcriptomes can be explored at the single-cell level, single-cell profiling of proteomes is not yet established. Here we describe new single-molecule protein sequencing and identification technologies alongside innovations in mass spectrometry that will eventually enable broad sequence coverage in single-cell profiling. These technologies will in turn facilitate biological discovery and open new avenues for ultrasensitive disease diagnostics.

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Single-molecule FRET is a versatile tool to study nucleic acids and proteins at the nanometer scale. However, currently, only a couple of FRET pairs can be reliably measured on a single object, which makes it difficult to apply single-molecule FRET for structural analysis of biomolecules. Here, we present an approach that allows for the determination of multiple distances between FRET pairs in a single object. We use programmable, transient binding between short DNA strands to resolve the FRET efficiency of multiple fluorophore pairs. By allowing only a single FRET pair to be formed at a time, we can determine the pair distance with subnanometer precision. The distance between other pairs are determined by sequentially exchanging DNA strands. We name this multiplexing approach FRET X for FRET via DNA eXchange. Our FRET X technology will be a tool for the high-resolution analysis of biomolecules and nanostructures.

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Förster resonance energy transfer (FRET) is a useful phenomenon in biomolecular investigations, as it can be leveraged for nanoscale measurements. The optical signals produced by such experiments can be analyzed by fitting a statistical model. Several software tools exist to fit such models in an unsupervised manner but lack the flexibility to adapt to different experimental setups and require local installations. Here, we propose to fit models to optical signals more intuitively by adopting a semisupervised approach, in which the user interactively guides the model to fit a given data set, and introduce FRETboard, a web tool that allows users to provide such guidance. We show that our approach is able to closely reproduce ground truth FRET statistics in a wide range of simulated single-molecule scenarios and correctly estimate parameters for up to 11 states. On in vitro data, we retrieve parameters identical to those obtained by laborious manual classification in a fraction of the required time. Moreover, we designed FRETboard to be easily extendable to other models, allowing it to adapt to future developments in FRET measurement and analysis.

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DNA cyclization assay together with single-molecule FRET was employed to monitor protein-mediated bending of a short dsDNA (~ 100 bp). This method provides a simple and easy way to monitor the structural change of DNA in real-time without necessitating prior knowledge of the molecular structures for the optimal dye-labeling. This assay was applied to study how Anabaena sensory rhodopsin transducer (ASRT) facilitates loop formation of DNA as a possible mechanism for gene regulation. The ASRT-induced DNA looping was maximized at 50 mM of Na⁺, while Mg²⁺ also played an essential role in the loop formation.

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Fueled by ATP hydrolysis in N-ethylmaleimide sensitive factor (NSF), the 20S complex disassembles rigid SNARE (soluble NSF attachment protein receptor) complexes in single unraveling step. This global disassembly distinguishes NSF from other molecular motors that make incremental and processive motions, but the molecular underpinnings of its remarkable energy efficiency remain largely unknown. Using multiple single-molecule methods, we found remarkable cooperativity in mechanical connection between NSF and the SNARE complex, which prevents dysfunctional 20S complexes that consume ATP without productive disassembly. We also constructed ATP hydrolysis cycle of the 20S complex, in which NSF largely shows randomness in ATP binding but switches to perfect ATP hydrolysis synchronization to induce global SNARE disassembly, minimizing ATP hydrolysis by non-20S complex-forming NSF molecules. These two mechanisms work in concert to concentrate ATP consumption into functional 20S complexes, suggesting evolutionary adaptations by the 20S complex to the energetically expensive mechanical task of SNARE complex disassembly.@en

The three-dimensional organization of DNA is increasingly understood to play a decisive role in vital cellular processes. Many studies focus on the role of DNA-packaging proteins, crowding, and confinement in arranging chromatin, but structural information might also be directly encoded in bare DNA itself. Here, we visualize plectonemes (extended intertwined DNA structures formed upon supercoiling) on individual DNA molecules. Remarkably, our experiments show that the DNA sequence directly encodes the structure of supercoiled DNA by pinning plectonemes at specific sequences. We develop a physical model that predicts that sequence-dependent intrinsic curvature is the key determinant of pinning strength and demonstrate this simple model provides very good agreement with the data. Analysis of several prokaryotic genomes indicates that plectonemes localize directly upstream of promoters, which we experimentally confirm for selected promotor sequences. Our findings reveal a hidden code in the genome that helps to spatially organize the chromosomal DNA.

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Most everyday processes in life involve a necessity for an entity to locate its target. On a cellular level, many proteins have to find their target to perform their function. From gene-expression regulation to DNA repair to host defense, numerous nucleic acid-interacting proteins use distinct target search mechanisms. Several proteins achieve that with the help of short RNA strands known as guides. This review focuses on single-molecule advances studying the target search and recognition mechanism of Argonaute and CRISPR (clustered regularly interspaced short palindromic repeats) systems. We discuss different steps involved in search and recognition, from the initial complex prearrangement into the target-search competent state to the final proofreading steps. We focus on target search mechanisms that range from weak interactions, to one- and three-dimensional diffusion, to conformational proofreading. We compare the mechanisms of Argonaute and CRISPR with a well-studied target search system, RecA.

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At the core of homologous DNA repair, RecA catalyzes the strand exchange reaction. This process is initiated by a RecA loading protein, which nucleates clusters of RecA proteins on single-stranded DNA. Each cluster grows to cover the single-stranded DNA but may leave 1- to 2-nucleotide (nt) gaps between the clusters due to three different structural phases of the nucleoprotein filaments. It remains to be revealed how RecA proteins eliminate the gaps to make a seamless kilobase-long filament. We develop a single-molecule fluorescence assay to observe the novel internal dynamics of the RecA filament. We directly observe the structural phases of individual RecA filaments and find that RecA proteins move their positions along the substrate DNA to change the phase of the filament. This reorganization process, which is a prerequisite step for interjoining of two adjacent clusters, requires adenosine triphosphate hydrolysis and is tightly regulated by the recombination hotspot, Chi. Furthermore, RecA proteins recognize and self-align to a 3-nt-period sequence pattern of TGG. This sequence-dependent phase bias may help the RecA filament to maintain structural integrity within the kilobase-long filament for accurate homology search and strand exchange reaction.@en

Nucleosomes form the unit structure of the genome in eukaryotes, thereby constituting a fundamental tenet of chromatin biology. In canonical nucleosomes, DNA wraps around the histone octamer in a left-handed toroidal ramp. Here, in single-molecule magnetic tweezers studies of chaperone-assisted nucleosome assembly, we show that the handedness of the DNA wrapping around the nucleosome core is intrinsically ambidextrous, and depends on the pre-assembly supercoiling state of the DNA, i.e., it is not uniquely determined by the octameric histone core. Nucleosomes assembled onto negatively supercoiled DNA are found to exhibit a left-handed conformation, whereas assembly onto positively supercoiled DNA results in right-handed nucleosomes. This intrinsic flexibility to adopt both chiralities is observed both for canonical H3 nucleosomes, and for centromere-specific variant CENP-A nucleosomes. These data support recent advances suggesting an intrinsic adaptability of the nucleosome, and provide insights into how nucleosomes might rapidly re-assemble after cellular processes that generate positive supercoiling in vivo.

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DNA supercoiling crucially affects cellular processes such as DNA replication, gene expression, and chromatin organization. However, mechanistic understanding of DNA supercoiling and the related DNA-processing enzymes has remained limited, mainly due to the lack of convenient experimental tools to probe these phenomena. Here, we report a novel high-throughput single-molecule assay for real-time visualization of supercoiled DNA molecules, named ISD (Intercalation-induced Supercoiling of DNA). We use an intercalating dye to induce supercoiling of surface-attached DNA molecules as well as to visualize coiled-loop structures (i.e., plectonemes) formed on DNA. The technique is solely based on epifluorescence microscopy and requires no mechanical manipulation of the DNA molecules. This new assay allows to track positions and sizes of individual plectonemes and characterize their position-dependent dynamics such as nucleation, termination, and diffusion. We describe the ISD technique and demonstrate its potential by establishing that plectonemes are pinned to a local 10-nucleotide long mispaired sequence along a double-stranded DNA molecule.

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Centromere-specific nucleosomes are a central feature of the kinetochore complex during mitosis, in which microtubules exert pulling and pushing forces upon the centromere. CENP-A nucleosomes have been assumed to be structurally unique, thereby providing resilience under tension relative to their H3 canonical counterparts. Here, we directly test this hypothesis by subjecting CENP-A and H3 octameric nucleosomes, assembled on random or on centromeric DNA sequences, to varying amounts of applied force by using single-molecule magnetic tweezers. We monitor individual disassembly events of CENP-A and H3 nucleosomes. Regardless of the DNA sequence, the force-mediated disassembly experiments for CENP-A and H3 nucleosomes demonstrate similar rupture forces, life time residency and disassembly steps. From these experiments, we conclude that CENP-A does not, by itself, contribute unique structural features to the nucleosome that lead to a significant resistance against force-mediated disruption. The data present insights into the mechanistic basis for how CENP-A nucleosomes might contribute to the structural foundation of the centromere in vivo.@en