Gary Pielak, a professor of chemistry, biochemistry, and biophysics at the University of North Carolina, Chapel Hill, uses a very different physical process to study protein dynamics in vivo. In fact, Pielak's technique might seem more at home in a chemistry building than a biology lab: nuclear magnetic resonance.
For years, Pielak has been interested in the impact macromolecular crowding has on protein stability. As an assistant professor studying proteins in dilute buffers in a test tube, a colleague pointed out that his model system probably did not accurately reflect life inside a cell. “This bugged me,” he recalls. He's spent the past 15 years trying to resolve the problem.
Inside a cell macromolecular concentrations are on the order of 300 gm/L or higher — three times more concentrated than an egg white and certainly more concentrated than the buffers Pielak previously used. Researchers have assumed that such a high volume occupancy, in and of itself, can constrain reactions, motion, and folding in vivo, simply by raising local concentrations, restricting motion, and limiting available space. “That is going to favor folded, collapsed states over unfolded ones, because unfolded structures take more volume,” he says.
That was the theory anyway; but Pielak wanted to know if it was true. To find out, he and his team created a mutant, isotopically labeled protein that was mostly unfolded in vitro, except in the presence of salt. They then asked whether macromolecular crowding in bacterial cells would push this protein's equilibrium toward a more folded state.
To figure that out, the team expressed isotopically (15N and 19F) labeled protein at high levels in bacteria — so high, in fact, that he calls his technique “in cell NMR” rather than in vivo, as the expression far exceeds normal levels. The cell slurry was then placed in an NMR tube — Pielak compares the mixture to “a melted vanilla milkshake”— and bombarded with radio frequency radiation to assess the chemical environment of the protein backbone, such as how exposed it is to buffer.
To their surprise, the team found that the protein was no more folded in vivo than in vitro, even under “hyperosmotic” conditions that forced the cells to take up more salt. “This idea that crowding will always stabilize proteins, is just not true,” Pielak concludes. Instead, he suggests a competition could be occurring between the influence of crowding, which would tend to favor the folded protein, and non-specific interactions between the unfolded protein and other macromolecules in the cell, which favor an unfolded form.
“Although there is an excluded volume effect [caused by crowding], the non-specific interactions favor the denatured state, and they win,” he says.
Several other recently developed methods also have the potential to shed new light on the ways in which proteins fold in cells. In February 2011, for instance, Henry Chapman of the Center for Free-Electron Laser Science at the University of Hamburg, Germany, described a technique called “femtosecond X-ray protein nanocrystallography,” a high-speed approach that bombards a moving stream of nanoscale protein crystals smaller than one-micron in size with intense bursts of X-ray energy (each pulse is one billion times brighter than synchrotron radiation). Chapman and his colleagues used three million diffraction patterns generated in this manner to produce an 8-Angstrom structure of photosystem-I.
Ultimately, the goal is to tune the laser to image individual proteins, says Chapman, though he admits “it's by no means a sure thing.” For one, focusing the laser to so small a spot requires new engineering. And it isn't clear how matter will behave under such intense radiation. If it works, though, it could make in vitro folding studies possible, he says, assuming it were possible to synchronize their motion in some way.
Another potentially useful technology: photoactivatable probes. Jennifer Lippincott-Schwartz, a Distinguished NIH Investigator at the National Institute of Child Health & Human Development, has developed photoactivatable fluorescent proteins — molecules that are “dark” unless they first are activated by a pulse of light — and applied them to a superresolution microscopy technique called PALM.
Photoactivatable FRET probes, she says, could be used as FRET acceptors in, for instance, pulse-chase experiments that track specific molecular populations over time. Or they could be combined with PALM to localize particular complexes with nanometer-precision.
Of course, it's not the technique that matters so much as the insight it provides. And according to Weninger, the problem right now is that most of the insights structural biology has so far provided have been based on unmodified bacterially expressed proteins studied in isolation. “We want to look at how the structure of proteins is related to function in a fully functioning living system,” he says. “[But] those kinds of complicated interactions are difficult to reproduce in vitro.”