Wesley Philip Wong

Molecules that modulate protein-protein interactions, including through ternary complex formation, have been established as key therapeutics. Yet they can be challenging to discover, as standard screening approaches often lack throughput or fail to directly assay multibody interactions. We addressed this by combining DNA nanoswitch–based selection of nucleotide-encoded libraries with next-generation sequencing to create FIND (Functional Interaction–based Nanoswitch Discovery), which enables ultrahigh-throughput discovery of compounds that induce or increase protein-protein interactions. We demonstrate FIND by discovering nanobodies that stabilize the interaction between angiotensin-converting enzyme 2 and the receptor-binding domain of severe acute respiratory syndrome coronavirus 2 and by discovering agonists of peroxisome proliferator–activated receptor gamma from a DNA-barcoded small-molecule library. By enabling rapid discovery of molecular modulators, FIND could lead to new therapeutics across many conditions.

The emergence of a highly contagious coronavirus in 2019 led to an unprecedented need for large-scale diagnostic testing. The associated challenges, including reagent shortages, cost, deployment delays, and turnaround time, have all highlighted the need for an alternative suite of low-cost tests. Here, we demonstrate a test for SARS-CoV-2 RNA that provides direct detection of viral RNA and eliminates the need for costly enzymes. We employ DNA nanoswitches that respond to segments of the viral RNA by a change in shape that can be read by gel electrophoresis. A multitargeting approach samples 120 different viral regions to improve the limit of detection and provide robust detection of viral variants. We applied our approach to a cohort of clinical samples, positively identifying a subset of samples with high viral loads. Since our method directly detects multiple regions of viral RNA without amplification, it eliminates the risk of amplicon contamination and renders the method less susceptible to false positives. This tool can benefit diagnostic options for COVID-19 and future emerging outbreaks, providing a third option between amplification-based RNA detection and protein antigen detection. Ultimately, we believe that this tool can be adapted both for low-resource onsite testing as well as for monitoring viral loads in recovering patients.

Single-molecule techniques have transformed biological research by enabling direct observation and manipulation of individual molecules. These methods overcome ensemble averaging inherent in bulk measurements and facilitate studies under physiological stresses and out-of-equilibrium conditions. They have provided valuable insights into diverse biological processes, from stepping mechanisms of molecular motors to mechanical properties of biomolecules to the dynamic strength of intermolecular bonds. Advances in multiplexed and high-throughput single-molecule force spectroscopy methods are improving throughput, capabilities, and accessibility. In this review, we detail the evolution of multiplexed force spectroscopy technologies, highlighting key advances in instrumentation, molecular engineering, and analytical techniques. We discuss diverse applications spanning molecular biophysics, biomolecular sensing, proteomics, and cellular mechanobiology. Finally, we explore ongoing challenges and future opportunities and highlight how the impact of multiplexed single-molecule force spectroscopy can continue to grow through further developments in novel instrumentation, chemical tools, and innovative applications.

Molecular patterning at single‐molecule resolution could significantly advance biomolecular analysis and the engineering of functional surfaces by enabling precise control over the spatial arrangement and mechanical accessibility of individual biomolecules. Such control is particularly valuable for multiplexed single‐molecule assays, which reveal molecular mechanisms, non‐equilibrium behavior, and nanoscale mechanical properties through the application of mechanical force. Standard surface functionalization methods, however, often lack sufficient precision, programmability, or accessibility, resulting in random or sparse biomolecular arrangements. To address this, we have developed a light‐guided surface patterning method that can covalently organize oligonucleotides (oligos) without the need for lithographic equipment. Oligos with 3‐cyanovinylcarbazole (CNVK) nucleoside are crosslinked in precise spatial arrangements defined by UV patterns projected through a digital micromirror device (DMD), with beads arranged accordingly. We demonstrate compatibility with established single‐molecule methods by performing single‐molecule force spectroscopy experiments on patterned coverslips, using magnetic tweezers and hydrodynamic‐based approaches. Our light‐guided method provides a scalable and accessible platform for biomolecular patterning that allows precise control over molecular identity and spatial positioning, enabling high‐throughput single‐molecule manipulation and mechanical characterization.

Understanding the three-dimensional structure and mechanical response of biomolecules is key to uncovering their molecular mechanisms, particularly in contexts where force plays a regulatory role. Structural methods such as X-ray crystallography, Cryo-electron microscopy, and Nuclear Magnetic Resonance (NMR) spectroscopy provide high-resolution conformational data, while single-molecule force spectroscopy reveals mechanical properties—but these approaches are rarely integrated. A more comprehensive understanding of structure-function relationships, including nonequilibrium conformations and transitions under force, calls for methods capable of simultaneously resolving structural and mechanical properties at the single-molecule level. To meet this need, we present a DNA nanoswitch calipers platform capable of both measuring multiple intramolecular distances and mechanically unfolding individual biomolecules along defined axes. Using human telomeric DNA G-quadruplexes as a model system, we mapped distances between labeled sites to distinguish conformational states and performed directional unfolding to characterize mechanical stability along defined axes. This integrative approach revealed subtle conformational and mechanical differences, showcasing DNA nanoswitch calipers as a modular, broadly applicable approach for mechanostructural analysis of complex biomolecular systems.

Molecular interactions between receptors and ligands govern critical biological processes, from immune surveillance and T-cell activation to tissue development. However, current techniques for studying binding avidity often sacrifice throughput or precision. We introduce a high-throughput method for quantifying molecular and cellular binding kinetics using a centrifuge force microscope (CFM)a compact imaging system integrated into a benchtop centrifuge. The CFM performs real-time force measurements on thousands of single cells in parallel, probing receptor–ligand interactions under controlled mechanical stress. To extend these capabilities, we developed a next-generation CFM with dual-channel fluorescence imaging that enables tracking of individual cell unbinding events. To demonstrate its utility, we profiled the binding mechanics of Bispecific T-cell Engager (BiTE) molecules, immunotherapeutic proteins that facilitate T-cell targeting of cancer cells. In cell–protein assays, we quantified the avidity of T and B cells interacting with BiTE-modified surfaces, revealing receptor-specific correlations between ligand concentration and bond strength. In cell–cell assays, we characterized BiTE-mediated adhesion between Jurkat and Nalm6 cells, demonstrating a time-dependent increase in avidity. By integrating force spectroscopy with fluorescence imaging, the CFM provides a high-throughput approach for investigating the mechanochemical principles underlying receptor-mediated interactions, with broad implications for biophysical chemistry, molecular recognition, and therapeutic development.

Understanding the three-dimensional structure and mechanical response of biomolecules is key to uncovering their molecular mechanisms, particularly in contexts where force plays a regulatory role. Structural methods such as X-ray crystallography, cryo-electron microscopy, and NMR spectroscopy provide high-resolution conformational data, while single-molecule force spectroscopy reveals mechanical properties—but these approaches are rarely integrated. A more comprehensive understanding of structure-function relationships, including non-equilibrium conformations and transitions under force, calls for methods capable of simultaneously resolving structural and mechanical properties at the single-molecule level. To meet this need, we present a DNA nanoswitch calipers platform capable of both measuring multiple intramolecular distances and mechanically unfolding individual biomolecules along defined axes. Using human telomeric DNA G-quadruplexes as a model system, we mapped distances between labeled sites to distinguish conformational states and performed directional unfolding to characterize mechanical stability along defined axes. This integrative approach revealed subtle conformational and mechanical differences, showcasing DNA nanoswitch calipers as a modular, broadly applicable approach for mechanostructural analysis of complex biomolecular systems.

Single-molecule force spectroscopy enables the detailed probing of molecular interactions, providing new insights into molecular mechanisms—yet studying biology “one molecule at a time” can lead to throughput challenges that limit applications. While multiplexed single-molecule assays can address these issues, suitable functionalization of surfaces is required, which remains a technical challenge—many commonly used approaches are constrained by random and sparse biomolecules arrangements, limiting programmability and throughput. An ideal anchoring method would enable (i) high surface densities to maximize throughput, (ii) precise control of spatial position and molecular identity for maximum control, (iii) covalent linking for high force application, and (iv) efficient patterning without the need for expensive facilities to maximize accessibility. To achieve these aims, we have developed a light-guided surface patterning method that can covalently organize oligonucleotides (oligos) without the need for lithographic equipment. Oligos with 3-Cyanovinylcarbazole (CNVK) nucleoside are crosslinked by UV patterns reflected through a digital micromirror device (DMD), with beads arranged accordingly. To demonstrate compatibility with established single-molecule methods, we performed single-molecule force spectroscopy experiments on patterned coverslips, using both magnetic tweezers and hydrodynamic-based systems. Our light-guided approach provides a scalable and accessible solution for biomolecular patterning that allows precise control over molecular identity and spatial positioning, enabling high-throughput measurements in single-molecule research.

Cell-cell binding, mediated by the physical interactions of receptors and their ligands, plays a fundamental role in immune processes such as immune surveillance and T-cell activation. However, current approaches for measuring cell avidity often lack either throughput or quantitative precision. Here, we introduce a high-throughput approach for quantifying cell binding lifetimes and strength using a centrifuge force microscope (CFM)—a compact microscope operating within a standard benchtop centrifuge. The CFM enables live monitoring of single-cell interactions under force, conducting thousands of force experiments in parallel. To facilitate the real-time study of live cell interactions, we developed a next-generation CFM with multichannel fluorescence imaging capabilities. This system accommodates measurements in two modes: cell-protein binding and cell-cell avidity assays. Using this system, we investigated immune-cell binding mediated by Bispecific T-cell Engager (BiTE) molecules, a novel immunotherapy designed to enhance immune-cell targeting of cancer cells. In cell-protein assays, we quantified T- and B-cell unbinding from BiTE-functionalized surfaces, revealing receptor-specific relationships between ligand concentration and binding strength. In cell-cell assays, we examined BiTE-mediated binding of T-cells to Nalm6 B-cells, a precursor leukemia cell line, uncovering a strong, time-dependent increase in BiTE-mediated immune-cell avidity. By integrating high-throughput and quantitative single-cell force analysis, the CFM provides new insights into the dynamic nature of immunological interactions under force, with broad implications for immunotherapy and cellular mechanics.

The ability to accurately map the 3D geometry of single-molecule complexes in trace samples is a challenging goal that would lead to new insights into molecular mechanics and provide an approach for single-molecule structural proteomics. To enable this, we have developed a high-resolution force spectroscopy method capable of measuring multiple distances between labeled sites in natively folded protein complexes. Our approach combines reconfigurable nanoscale devices, we call DNA nanoswitch calipers, with a force-based barcoding system to distinguish each measurement location. We demonstrate our approach by reconstructing the tetrahedral geometry of biotin-binding sites in natively folded streptavidin, with 1.5-2.5 Å agreement with previously reported structures.