Our research involves combining the tools and methods of physics and molecular biochemistry to investigate the response of single biological macromolecules to mechanical forces. In particular, we are interested in using force to probe the nanomechanics of structure formation and dissolution. "Folding," the formation of complex three-dimensional structures from linear biomolecular chains like nucleic acids and proteins, is one of the most important examples of self-assembly that we know of: biological molecules must fold into the correct shape in order to function correctly.
Single-molecule measurements of unfolding and refolding under mechanical tension can offer important new insights into folding because they allow us to follow the progress of the folding reaction in individual molecules, with high spatial and temporal resolution. By measuring the extension of a molecule as a function of time at various forces, individual folding and unfolding events can be observed, the free energy and rates can be determined directly, intermediate states along the folding pathway can be observed, and the shape of the energy landscape governing the folding transition can even be deduced.
The main tool we use in this work is the optical trap, consisting of a tightly focused, intense laser beam which can hold on to micron-sized plastic beads in solution. An optical trap acts like a spring made out of light, allowing force to be applied to a molecule tethered to a bead by deflecting the bead from the center of the trap. The motion of the molecule is detected by collecting light scattered off the bead. State-of-the-art optical traps can detect motions of as little as 0.1 nm (the radius of a hydrogen atom!), providing an exceptionally precise tool for measuring folding trajectories. We are currently using optical traps to study the folding of simple nucleic acids and proteins, investingating the effects on the folding of changes in the molecular sequence and environmental conditions (buffer salt, pH, ...).