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DNA and RNA form the fundamental building blocks of the universal genetic code. They can form complex structures which interact with proteins, other nucleic acids, and even small regulatory molecules. Understanding the fundamental properties of these molecules gives us not only important physical insight into the physics of polymers, but also into the role their structures and interactions play in biological systems, from the simplest DNA-DNA interactions all the way up to their role in complex macromolecular machines like RNA polymerase and the ribosome. RNA can even play a role as an enzyme (so-called ribozymes) which can directly catalyze chemical reactions and regulate genetic expression.
In the Block Lab, we are capable of directly measuring and applying forces to nucleic acid polymers using optical trapping. This capability gives us the ability to study them from a unique perspective, one which is complementary to more traditional biochemical approaches. We have used this approach to probe the fundamental thermodynamics of DNA and RNA structures, as well as the biological function of riboswitches. We have also been involved in developing new technologies for combining fluorescence with optical trapping.
A DNA "dumbbell" trapped in a passive optical force clamp.
We have investigated the mechanical properties of nucleic acids by focusing in particular on hairpins. These structures consist of single strands of DNA or RNA whose ends are self-complementary, such that they loop back on themselves to form a duplex "stem" connected to a single-stranded loop (inset below). Hairpins not only provide a model system for studying DNA unzipping, but are also important in their own right (for example, they are a principal motif of secondary structure in RNA, and they play a role in gene transcription and regulation). The aim of our research is to understand, at a fundamental level, the factors that affect the stability of hairpins.
Dumbbell experimental geometry with DNA hairpin. From Woodside, et al., PNAS 2006.
In our first study of DNA hairpins (Woodside, Behnke-Parks, Larizadeh, Travers, Herschlag, and Block, PNAS 2006), we measured the kinetics and energetics of folding for a set of hairpins all having “pseudorandom” stem sequences, but having widely varying stem sizes, loop sizes and percentages of GC basepairs. This study introduced a simple model for the energy landscape of a folding hairpin, and found that the energy barrier between the folded and unfolded states corresponds to a distance 1-2 bp from the loop. In the second study (Woodside, Anthony, Behnke-Parks, Larizadeh, Herschlag, and Block, Science 2006), we turned to non-random (or “patterned”) hairpin stem sequences having mismatches or engineered blocks of GC and AT basepairs, to demonstrate that we could manipulate the energy landscape by introducing intermediate states or changing the height or location of the energy barrier. To further test our model, measured the full energy landscapes of four hairpin sequences by measuring histograms of the end-to-end extensions of these hairpins over time, and then removing noise from the histograms by deconvolution.
Riboswitches are mRNA elements that regulate gene expression through ligand-induced conformational changes. They typically lie in the 5’ untranslated region of some genes and consist of a ligand binding aptamer domain followed by an expression platform. Aptamer conformation affects the structure adopted by the expression platform (e.g., the formation or absence of a terminator hairpin) and thus the gene’s expression.
Among the simplest riboswitches are those involved in purine metabolism, which contain aptamers with a three helix junction. We have studied the aptamer of one such purine riboswitch, the pbuE adenine riboswitch.
pbuE adenine riboswitch aptamer. ‘A’ indicates
binding of the aptamer’s ligand, adenine.
Using our single molecule dumbbell assay we are able to effectively hold the RNA of interest between two beads and exert force on it in order to probe aptamer structure. In general, we measure extension change during folding and unfolding of the aptamer and relate extension changes to numbers of nucleotides and ultimately to specific aptamer structures.
Dual-trap optical tweezers assay showing the experimental geometry with stalled RNAP (green) and nascent RNA transcript (red) containing the riboswitch aptamer sequence.
As an example, the figure below shows force-extension curves while unfolding the aptamer. In the presence of adenine (the aptamer’s ligand), the RNA is often ligand bound and stabilized; it only rips to the unfolded state at high force.
Force-extension curves showing aptamer unfolding. Without adenine, two events are seen (black), corresponding to the unfolding of hairpins P2 and P3 (inset). With adenine bound to the aptamer, large unfolding events are observed (blue), sometimes involving an intermediate state (red), that correspond to opening of the entire aptamer structure.
We have extracted energies, distances, and rate constants describing aptamer folding by using force to study aptamer kinetics and equilibrium folding transitions. The adenine-induced stabilization of the closing aptamer helix, P1, was described and a quantitative energy landscape for riboswitch aptamer folding was created, dissecting the secondary and tertiary folding events of this RNA structure.
To learn more about this work see Greenleaf, et al. Science, 2008.
Combined Optical Trapping and Fluorescence Resonance Energy Transfer
Our lab has recently developed new technologies which combine optical trapping with fluorescence (Lang et al., Nat. Methods 2004, Lang et al. J. Biology 2003), to extend the detection capabilities of the instrument. One such experiment examined the force required to separate short segments of double-stranded DNA. These measurements included the additional element of fluorescent reporting of the DNA unzipping: single fluorophores were attached to each of the two strands of DNA, and their fluorescence was monitored at the same time as force was applied to the DNA with the optical tweezers (figure on left, below). A typical measurement is shown below on the right. At first, the fluorescence (blue trace) is quenched because the fluorophores are close to each other. As we start to pull on the DNA, the force (red trace) increases until the DNA unzips; at this point, the two strands separate, and the fluorescence increases. This technique of combined optical trapping and single-molecule fluorescence should prove to be a powerful and quite general tool for studying single-molecule systems, because it allows us to pinpoint the location of structural changes and to determine the relative timing of events.
The combination of single molecule fluorescence and optical tweezers in a hairpin unzipping experiment. The addition of the fluorescence signal allows us to better locate, both in space and time, a known structural event relative to an event observed with the optical tweezers (Figure from Lang et al. J. Biology 2003).
Another recent development is the ability to perform fluorescence resonance energy transfer (FRET) measurements concurrently with optical trapping. Our instrument utilizes a dual trap configuration with fluorescence illumination between the traps. In the example below, a DNA hairpin is held between the traps (left), with fluorescent labels at the base of the hairpin. The hairpin is closed under low force, and opens under high force, with a rip signal evident in the force-vs-extension curve (black). As the hairpin opens, the fluorescence signals change (red and green), and a drop in FRET occurs (blue), due to the sudden increase in extension. We hope to apply this technology to the study of nucleic acid structure, by observing internal structural changes of RNA due to externally applied forces.