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Single-Molecule Biophysics

Single-Molecule Studies of DNA, RNA, and Protein

The study of single molecules of DNA, RNA, and protein is revolutionizing biophysics. To investigate the structure and function of selected biomolecules in all three categories, the Thomas Perkins group has developed and is using precision single-molecule biophysics techniques such as atomic force microscopy (AFM), single-molecule force spectroscopy, and optical tweezers, which are tiny micro-sized traps created by a tightly focused laser beam. For the latter technique, the group uses an optical-trapping assay it developed with Ångstrom-scale stability and resolution in three dimensions.

The group uses the three techniques to study the folding and unfolding of proteins in vitro with the goal of better understanding their function in living cells. It recently began an AFM study to image membrane proteins and pull on them with well-defined forces. The goal is to understand the folding and unfolding of these biomolecules in native lipid (fat) bilayers rather studying them as part of solubilized membranes. The group wants to better understand how the structure and behavior of these proteins affect their function. In related work, the group is using AFM to characterize mutants of bacterial rhodopsin, a light-sensitive protein.

The Perkins lab uses high-precision optical tweezers to investigate the mechanical properties of DNA, dye-DNA interactions, molecular motors that travel along strands of DNA, force standards based on biological molecules, as well as the folding and unfolding of RNA. Recent force-microscopy studies of dye-DNA interactions have revealed important dynamics of dye molecules that bind to and insert themselves into strands of DNA. The molecules, called intercalators, can be used to fluorescently label DNA and as drugs in cancer therapy.
 
The Perkins group has also used force microscopy to stretch DNA in the presence and absence of dye to study the effects of DNA binding and intercalation. This work has shown that binding/unbinding and intercalation/de-intercalation are distinct processes that occur on different time scales. As part of the work on DNA stretching, the group was able to show that peeling (i.e., the generation of single-stranded- (ss-) DNA from nicks or free ends) is not required for the 70% DNA overstretching observed in a narrow force window at 65 pN.
 
In related work on nucleic acids, the group is using force microscopy to investigate the energetics and folding pathways of riboswitches. Riboswitches are regulatory portions of a messenger RNA molecule that bind to small molecules. This binding causes a change in the production of proteins encoded by the mRNA molecule.

The group also investigates DNA protein motors, which bind to DNA and move along it. The researchers are currently exploring the process by which these proteins change chemical energy into physical motion. In support of this effort, the group has developed an optical-trapping microscope capable of resolving the smallest known step of a molecular motor, the 1-base pair (0.34 nm) step of along DNA. The group is now applying this technology to the study of DNA-based molecular motors and transcription factors, as well as the folding and unfolding kinetics of RNA and RNA-protein complexes.

New Nano Microscopy Zooms in on Membrane Proteins

The Tom Perkins and Markus Raschke groups recently demonstrated a new infrared (IR) imaging system with a spatial resolution and a chemical sensitivity of just a few membrane proteins. In their initial experiment, the researchers used the tip of an atomic force microscope (AFM) as an antenna for the IR light. The antenna focused the light onto a sample containing a bacterial protein called bacteriorhodopsin (bR). The AFM-tip antenna also helped capture the IR signal emitted by the bR protein and send it back to a detector for identification and location. The new antenna works much like a cell phone antenna except that it communicates with protein molecules.

The AFM-tip antenna works with an IR nano microscope called s-SNOM (scattering scanning near field optical microscope). The combination made it possible to identify the bR protein with a spatial recolution of 20 nm, or the length of 2–3 bR molecules. The researchers were also able to acquite IR spectra of just a handful of protein molecules rather than the approximately 10,000 molecules required for obtaining good spectra with an ordinary IR microscope. The success of this work has opened up imaging of biological and chemical structures 5000 times smaller than the diameter of a human hair.

The Raschke group is working on improving the new imaging method, with the goal of resolving single molecules, including some in liquid. For its part, the Perkins group continues its quest to better understand membrane proteins, which are the target of 50% of all current and future drugs. The two groups plan to continue their collaboration with the goal of being able to probe and understand the structures and functions of the constituents of living cells—in real time and under realistic conditions for life.

Single-Molecule Microscopy and Kinetics

The David Nesbitt group’s research into the biophysics of biomolecule folding began in the early 2000s with studies of the kinetics of folding of single RNA molecules in solution. Over the next decade, the group investigated the temperature dependence of RNA folding and unfolding as well as the influence of magnesium (Mg2+) and sodium (Na+) ions on conformation changes. The researchers found that both Mg2+ and Na+ ions neutralized negative charges that moved closer to one another in folded RNA pieces, a process that facilitated folding. Higher concentrations of these two ions also made the energy balance of folding more favorable.

During more than a decade of detailed biophysical analysis of RNA folding, the researchers assumed that what they discovered about single RNA-molecule folding in solution would translate to an understanding of RNA folding dynamics inside living cells. However, the researchers realized that conditions inside living cells are very different from carefully constructed in vitro experiments in the laboratory. In cells, proteins, nucleic acids (DNA and RNA), and other cellular materials take up as much as 80 % of the available volume—creating an extremely crowded environment.

Crowding changes the energy balance of folding and unfolding RNA because it disproportionately slows the process of unfolding. This effect makes it much more challenging to extrapolate results obtained on RNA folding in vitro to what goes on inside living cells. As a result, the Nesbitt group has undertaken a long-term effort to better understand molecular crowding and its impact on the folding of RNA and proteins.

The group also investigates the formation of viral protein coats, or capsids, that surround infectious RNA molecules. The capsids are made from a single long protein coded for by viral RNA and produced by the host cell. The capsid protein folds into protein fragments that come together to form the capsid, a structure that resembles a soccer ball. The capsid encloses new viral RNA and is ejected from the host cell, where it can spread the viral infection to other cells.

The researchers are interested in several steps in this complex process. They are looking at early RNA-RNA interactions that form larger strands of RNA coding for the capsid protein. They are also investigating the RNA-protein interactions that lead to the formation of a new, infectious virus particle. For these studies, the group is investigating a member of the tobacco mosaic virus family and human immunodeficiency virus (HIV). In the HIV investigations, the researchers study how HIV knows to stop replicating itself and export new virus particles to another organism. In studies on both viruses, they have discovered that the infectious RNA molecule is capable of “self recognizing” sequences in different RNA molecules needed to create the capsid protein. Viral RNA also appears to trigger the final stage of viral capsid formation.

In related work, the researchers are looking at what happens at the level of 5–9 base pairs during DNA hybridization. They are particularly interested in how hybridization starts from “fraying ends” of DNA. They want to understand (at the molecular level) what controls and influences hybridization rates and better understand the energetics of the process.

The group also wants to better understand (at the molecular level) the processes involved in the denaturation and reformation of cellular proteins. To do this, the researchers have recently begun in vitro studies of the action of urea, a natural denaturant of proteins. Urea can unpeel both proteins and nucleic acids. The goal is to understand the process of a protein breaking apart at the single-molecule level.

The group has recently undertaken single-molecule studies of human telomerase, an RNA-containing enzyme. Telomerase has been implicated in both aging and cancer because it sits on DNA chromosomes and is critical to the initiation of DNA replication. In essence, telomerase creates a new landing pad for enzymes when they copy strands of DNA.

Without telomerase, cells rapidly age and die because they lose the ability to replicate DNA. For example, telomerase mutations cause diskeratosis congenita, a disease that appears in children between 5 and 15 years old and causes symptoms of premature aging, resulting in shortened life spans. Telomerase is also implicated in the proliferation of cancer cells because it is required for repeated DNA replication and cell division.

A better understanding of the biophysics of telomerase activity may lead to improved cancer therapies. To aid in this understanding, the Nesbitt group is investigating the folding kinetics of a double-looped RNA segment in telomerase known as the telomerase pseudoknot. The formation of this pseudo knot is essential for normal telomerase activity.