Single-molecule detection has opened vast avenues to investigate aspects of biological systems that are inaccessible by any other technique. However, most of the single-molecule studies have been limited to isolated single protein, RNA, and DNA molecules, yet these molecules do not function in isolation in the cell. To better emulate the cellular environment, we need to study more-complex systems with many components.
Research in my lab is focused on pushing the limits of single-molecule detection methods to study biological systems as a complex. To do this, we develop state-of-the-art techniques (e.g., multicolor fluorescence, super-resolution imaging, combined force and fluorescence spectroscopy, vesicular encapsulation, quantitative fluorescence labeling strategies). These techniques are enabling us to test diverse protein–nucleic acid and protein-protein complexes and the mechanistic basis of their interactions and functions—both in vitro and in vivo—at an unprecedented spatial and temporal resolution.
Conformational Dynamics of Nucleic Acids
The elastic model of DNA duplex dynamics predicts high bending stiffness in DNA polymers less than 150 base pairs (bp) in length. We recently developed a fluorescence resonance energy transfer (FRET)-based cyclization assay to explain the apparent bendability of DNA <100 bp observed in many cellular processes, such as gene expression regulation, packaging in viral capsids, and nucleosome formation. We can determine the effect of DNA length on the rates and variation of DNA looping. The DNA cyclization assay allows us to see the effect of various DNA modifications, defects, and proteins on DNA flexibility. With respect to RNA dynamics, we have used FRET measurements to understand folding properties of guanine-dependent purine riboswitches (in collaboration with Scott Silverman [University of Illinois at Urbana-Champaign]). We are also utilizing three-color FRET strategies and MD-Gō simulations (in collaboration with Sarah Woodson [Johns Hopkins University] and Zaida Luthey-Schulten [UIUC]) to understand the sequential events in ribosome assembly. Our studies of ribosomal protein S4 binding to Escherichia coli 16S rRNA have captured the cooperativity in simultaneous binding and folding events and shed more light on protein-RNA interactions during the ribosome assembly process.
Single-Molecule Study of Nucleic Acid–Binding Proteins
Helicases—the DNA/RNA-unwinding enzymes—and their enzymatic activities are associated with virtually all cellular processes involving nucleic acids and constitute an important aspect of our research. Helicases are found in all three kingdoms of life and are extremely numerous: an estimated 1–2 percent of eukaryotic genes encode helicases. Several severe human genetic diseases have been linked to mutations in these proteins. Using our small-molecule FRET approaches, we have gained insights into the functions of different families of helicases (e.g., Rep, NS3, UvrD, PcrA) and their role during DNA replication (e.g., T7). For example, we discovered that PcrA helicase reels in DNA in single base steps, forming a DNA loop and at the same time efficiently removing other proteins bound to the DNA.
We also discovered that NS3 helicase from a human pathogen, hepatitis C virus, uses a spring-loaded mechanism to unwind DNA. Furthermore, we determined that a "priming loop" formed during double-stranded DNA unwinding by T7 helicase enables coordination of leading- and lagging-strand synthesis during DNA replication. With respect to other DNA-binding proteins, we have used three-color FRET strategy to show that the sliding mechanism of RecA filament is crucial for fast homology search during DNA repair and recombination.
To elucidate the functional behavior of DNA-binding proteins in more detail, we have recently developed a next generation of single-molecule instrumentation combining ultrahigh-resolution optical traps, capable of detecting motions at the subnanometer scale, and single-molecule fluorescence detection. The instrument uses a high-speed interlacing scheme to combine dual optical traps with a confocal fluorescence microscope. This instrument will enable us to make direct correlation, for the first time, between conformational states and unwinding steps of helicases. In addition, we have recently built an instrument combining total internal reflection fluorescence microscopy with optical tweezers (fleezers) so that we can track movements of fluorescently labeled molecules on a stretched single-stranded DNA (ssDNA). This platform will allow us to probe the consequences of an encounter between a helicase and other DNA-bound proteins, which are difficult to probe using FRET alone. Insights into mechanisms of single-stranded DNA-binding protein (SSB) sliding on ssDNA are an example. We are expanding fleezer technology to study nucleosome conformational dynamics, particularly with respect to DNA wrapping and unwrapping on the histone octamers.
Visualizing Protein-Protein and Protein–Nucleic Acid Interactions
While we are taking advantage of single-molecule techniques to study biological molecules in vitro, we are also interested in understanding biological processes in their physiological context. We have developed a single-molecule pull-down (SiMPull) assay that combines the principles of a conventional immunoprecipitation assay with single-molecule fluorescence microscopy to enable direct visualization of individual cellular protein complexes from a variety of sources. We have utilized this method for diverse complex systems to detect rare subpopulations of protein complexes and determine their stoichiometric composition in their cellular environment (in collaborations with Peter Cresswell [HHMI, Yale School of Medicine], Supriya Prasanth [University of Illinois at Urbana-Champaign], and Jie Chen [UIUC]).
For visualization within a cell, we have constructed a two-color three-dimensional (3D) super-resolution optical microscope. Because living cells house many nanometer-sized molecules that are densely packed into assemblies and networks, traditional light microscopes, with their low resolving power, are insufficient. Our super-resolution fluorescence imaging achieves spatial resolution of 50 nm in all three dimensions; this allows us to spatially localize and rigorously investigate protein-protein, protein-RNA, and protein-DNA interactions inside cells. We are interested in protein-protein interactions during DNA repair processes in bacterial cells. In particular, we are trying to understand the bacterial SOS response through cellular colocalization of DNA repair proteins and their time-dependent interactions during the repair process.
Imaging the Antiviral Immune Response
We are also interested in the function of RIG-I and other RIG-I-like receptors (RLRs), cytoplasmic sensors that recognize pathogen-associated molecular patterns of viral RNA to initiate the innate immune response. We have shown the ATP-dependent translocation of RIG-I on double-stranded RNA (in collaboration with Karl-Peter Hopfner [University of Munich]), but the significance of the translocation activity remains unclear. To this end, we are using SiMPull, fluorescence in situ hybridization (FISH), and site-specific and unique protein-labeling techniques to understand the signaling and detection mechanisms of RLRs on the cellular level and on the single-molecule level. Through these techniques, we hope to discover important signaling partners of these RLRs, understand how this signal is transmitted throughout the cell in the presence of viral infections, and observe the dynamics of viral-like RNA detection.
Mechanical Perturbation and Response of Biological Systems
To study force-dependent cellular function, we are developing probes to target a variety of cellular receptors that allow us to sense forces either by FRET sensing or Rupture sensing, wherein we observe rupture of a sensor calibrated to rupture at a predefined force. Our fleezers instrument has enabled FRET versus force calibration at a much greater force and distance sensitivity. We have demonstrated FRET-force sensing by inserting an elastic peptide between the TFP-Venus fluorescent protein FRET pair. By placing this sensor in between the Vt and Vh domains of the protein vinculin, we have been able to achieve quantification of real-time focal adhesion formation by vinculin in live cells (in collaboration with Martin Schwartz [University of Virginia], Christopher Chen [University of Pennsylvania] and Stephen Sligar [UIUC]). We are also interested in understanding molecular forces that regulate cell surface receptor activation on the single-cell level. We are applying Rupture sensing to study long-term effects due to force intervention in cell–extracellular matrix (ECM) interaction. To understand the collective cell-adhesion process, we are using Rupture sensors based on DNA to measure forces experienced by single integrin-RGD complexes for different cell types.
We have recently developed a site-specific, fast, quantitative, and highly efficient protein-labeling protocol (in collaboration with Isaac Cann [UIUC]) based on genetic incorporation of aldehyde tags onto proteins. This method specifically labels the protein of interest in mild conditions in cellular extracts while preserving its biological function. We have also identified a library of bright and long-lasting fluorescent proteins (FPs) better suited for studying many biological processes that suffer because of the poor photostability of traditional FPs in single-molecule assays.
Another effort in our lab is aimed at breaking the concentration restrictions on single-molecule fluorescence methods, which are currently limited to low nanomolar concentrations. We have used vesicular encapsulation, which partially solves this issue by increasing the local effective concentration, to study SNARE-mediated fusion of single vesicles (in collaboration with Yeon-Kyun Shin [Iowa State University]). We are investigating new techniques to increase this limit to micromolar concentrations of labeled molecules, which will allow us to use single-molecule fluorescence methods to probe weak biological interactions.
Grants from the National Institutes of Health and the National Science Foundation provided partial support for these projects.
As of March 08, 2013