DNA Flow-Stretch Assays for Studies of Protein-DNA Interactions at the Single-Molecule Level

Protein-DNA interactions are the core of the cell’s molecular machinery. For a long time, conventional biochemical methods served as a powerful investigatory basis of protein-DNA interactions and target search mechanisms. Currently single-molecule (SM) techniques have emerged as a complementary tool for studying these interactions and have revealed plenty of previously obscured mechanistic details. In comparison to the traditional ones, SM methods allow direct monitoring of individual biomolecules. Therefore, SM methods reveal reactions that are otherwise hidden by the ensemble averaging observed in conventional bulk-type methods. SM biophysical techniques employing various nanobiotechnology methods for immobilization of studied molecules grant the possibility to monitor individual reaction trajectories of biomolecules. Next-generation in vitro SM biophysics approaches enabling high-throughput studies are characterized by much greater complexity than the ones developed previously. Currently, several high-throughput DNA flow-stretch assays have been published and have shown many benefits for mechanistic target search studies of various DNA-binding proteins, such as CRISPR-Cas, Argonaute, various ATP-fueled helicases and translocases, and others. This review focuses on SM techniques employing surface-immobilized and relatively long DNA molecules for studying protein-DNA interaction mechanisms.

PCR-free, label-free detection of sequence-specific DNA with single-molecule sensitivity using in vitro N-hybrid system in microfluidic drops

We propose a novel method that can detect DNA with high specificity at the single-molecule level by employing the in vitro N-hybrid strategy realized in sub-picoliter microfluidic drops. It detects target DNA based on the specific interactions of the target-encoded proteins with their partner molecules, and achieves single-molecule sensitivity via signal-transduction and signal-amplification during gene-expression processes in a sub-picoliter droplet, therefore effectively avoiding complicated procedures in labeling-based methods or biases and artifacts in PCR-based methods.

Mobility of nucleosomes is regulated by their binding partners

Chromatin accessibility is modulated in a variety of ways, both to create open and closed chromatin states which are critical for eukaryotic gene regulation. At the mechanistic single molecule level, how accessibility is regulated remains a fundamental question in the field. Here, we use single molecule tracking by high-speed atomic force microscopy to investigate this question using chromatin arrays and extend our findings into the nucleus. By high-speed atomic force microscopy, we tracked chromatin dynamics in real time and observed that the essential kinetochore protein CENP-C reduces the diffusion constant of CENP-A nucleosomes and the linker H1.5 protein restricts H3 nucleosome mobility. We subsequently interrogated how CENP-C modulates CENP-A chromatin dynamics in vivo. Overexpressing CENP-C resulted in reduced centromeric transcription and impaired loading of new CENP-A molecules. These data suggest a model in which inner kinetochore proteins are critically involved in modulating chromatin accessibility and consequently, noncoding transcription at human centromeres.

Modified Cytosines versus Cytosine in a DNA Polymerase: Retrieving Thermodynamic and Kinetic Constants at the Single Molecule Level

DNA methylation plays key roles in various areas, such as gene expression, regulation, epigenetics, and cancers. Since 5-methylcytosine (5mC) is commonly present in methylated DNA, characterizing the binding kinetics and...

A nanopore interface for high bandwidth DNA computing

DNA has emerged as a powerful substrate for programming information processing machines at the nanoscale. Among the DNA computing primitives used today, DNA strand displacement (DSD) is arguably the most popular, with DSD-based circuit applications ranging from disease diagnostics to molecular artificial neural networks. The outputs of DSD circuits are generally read using fluorescence spectroscopy. However, due to the spectral overlap of typical small-molecule fluorescent reporters, the number of unique outputs that can be detected in parallel is limited, requiring complex optical setups or spatial isolation of reactions to make output bandwidths scalable. Here, we present a multiplexable sequencing-free readout method that enables real-time, kinetic measurement of DSD circuit activity through highly parallel, direct detection of barcoded output strands using nanopore sensor array technology (Oxford Nanopore Technologies' MinION device). We show that engineered reporter probes can be detected and classified with high accuracy at the single-molecule level directly from raw nanopore signals using deep learning. We then demonstrate this method's utility in multiplexed detection of clinically relevant microRNA sequences. These results increase DSD output bandwidth by an order of magnitude over what is possible with fluorescence spectroscopy, laying the foundations for a new paradigm in DNA circuit readout and programmable multiplexed molecular diagnostics using portable nanopore devices.

Real-time, single-molecule observation of biomolecular interactions inside nanophotonic zero mode waveguides

Living cells rely on numerous protein-protein, RNA-protein and DNA-protein interactions for processes such as gene expression, biomolecular assembly, protein and RNA degradation. Single-molecule microscopy and spectroscopy are ideal tools for real-time observation and quantification of nucleic acids-protein and protein-protein interactions. One of the major drawbacks of conventional single-molecule imaging methods is low throughput. Methods such as sequencing by synthesis utilizing nanofabrication and single-molecule spectroscopy have brought high throughput into the realm of single-molecule biology. The Pacific Biosceinces RS2 sequencer utilizes sequencing by synthesis within nanophotonic zero mode waveguides. A number of years ago this instrument was unlocked by Pacific Biosciences for custom use by researchers allowing them to monitor biological interactions at the single-molecule level with high throughput. In this capability letter we demonstrate the use of the RS2 sequencer for real time observation of DNA-to-RNA transcription and RNA-protein interactions. We use a relatively complex model – transcription of structured ribosomal RNA from E. coli and interactions of ribosomal RNA with ribosomal proteins. We also show evidence of observation of transcriptional pausing without the application of an external force (as is required for single-molecule pausing studies using optical traps). Overall, in the unlocked, custom mode, the RS2 sequencer can be used to address a wide variety of biological assembly and interaction questions at the single-molecule level with high throughput. This instrument is available for use at the Center for Integrated Nanotechnologies Gateway located at Los Alamos National Laboratory.

Conformational heterogeneity of molecules physisorbed on a gold surface at room temperature

A quantitative single-molecule tip-enhanced Raman spectroscopy (TERS) study at room temperature remained a challenge due to the rapid structural dynamics of molecules exposed to air. Here, we demonstrate the hyperspectral TERS imaging of single or a few brilliant cresyl blue (BCB) molecules at room temperature, along with quantitative spectral analyses. Robust chemical imaging is enabled by the freeze-frame approach using a thin Al2O3 capping layer, which suppresses spectral diffusions and inhibits chemical reactions and contaminations in air. For the molecules resolved spatially in the TERS image, a clear Raman peak variation up to 7.5 cm-1 is observed, which cannot be found in molecular ensembles. From density functional theory-based quantitative analyses of the varied TERS peaks, we reveal the conformational heterogeneity at the single-molecule level. This work provides a facile way to investigate the single-molecule properties in interacting media, expanding the scope of single-molecule vibrational spectroscopy studies.

DNA fluctuations reveal the size and dynamics of topological domains

DNA supercoiling is a key regulatory mechanism that orchestrates DNA readout, recombination, and genome maintenance. DNA-binding proteins often mediate these processes by bringing two distant DNA sites together, thereby inducing (transient) topological domains. In order to understand the dynamics and molecular architecture of protein induced topological domains in DNA, quantitative and time-resolved approaches are required. Here we present a methodology to determine the size and dynamics of topological domains in supercoiled DNA in real-time and at the single molecule level. Our approach is based on quantifying the extension fluctuations -in addition to the mean extension- of supercoiled DNA in magnetic tweezers. Using a combination of high-speed magnetic tweezers experiments, Monte Carlo simulations, and analytical theory, we map out the dependence of DNA extension fluctuations as a function of supercoiling density and external force. We find that in the plectonemic regime the extension variance increases linearly with increasing supercoiling density and show how this enables us to determine the formation and size of topological domains. In addition, we demonstrate how transient (partial) dissociation of DNA bridging proteins results in dynamic sampling of different topological states, which allows us to deduce the torsional stiffness of the plectonemic state and the kinetics of protein-plectoneme interactions. We expect our approach to enable quantification of the dynamics and reaction pathways of DNA processing enzymes and motor proteins, in the context of physiologically relevant forces and supercoiling densities.