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Mechanical Studies of Single Molecules
Summary: Carlos Bustamante uses novel methods of single-molecule visualization, such as scanning force microscopy, to study the structure and function of nucleoprotein assemblies. His laboratory is developing methods of single-molecule manipulation, such as optical and magnetic tweezers, to characterize the elasticity of DNA, to induce the mechanical unfolding of individual protein and RNA molecules, and to investigate the machine-like behavior of molecular motors.
Since the publication of the classical studies of the Dutch scientist J. D. van der Waals in 1873, physical scientists have known that many—if not most—properties of matter can be rationalized by the strength and direction of the forces that molecules exert on each other. Even strictly macroscopic phenomena, such as the elasticity and the melting points of solids, the viscosity and boiling points of liquids, or the compressibility of gases, are macroscopic manifestations of the myriad of small interactions between molecules. Chemists have also known that chemical affinity results from the attractive interactions between chemical entities. In 1889, Svante Arrhenius proposed that reactions between molecular species follow pathways that involve the formation of some type of strained, largely unstable, high-energy transition state whose accessibility along the reaction coordinate controls the rate of the reaction.
Until very recently, chemists and biochemists have had to rely on bulk methods to investigate the properties of molecules and their reactions. During the last few years, however, the advent of novel methods of single-molecule manipulation have begun to offer researchers, for the first time, the opportunity to measure directly the forces holding together molecular structures, to measure the stresses and strains generated during the course of chemical and biochemical reactions, to exert external forces to alter the fate of these reactions, and to reveal the rules that govern the interconversion of mechanical and chemical energy in these reactions. This area of research can be rightly called mechanochemistry.
Biochemical processes as diverse as protein folding, DNA elasticity, the protein-induced bending of DNA, the stress-induced catalysis of enzymes, the mechanical properties of molecular motors, and even the ubiquitous processes of induced-fit molecular recognition are all examples in which stresses and strains develop in molecules as they move along a reaction coordinate. My group has played a central role in the development of this field of work.
Transcription by Yeast RNA Polymerase II
RNA polymerase II (RNAP II) is responsible for transcribing all mRNAs in eukaryotic cells in a highly regulated process that is conserved from yeast to human and that serves as a central control point for cellular function. The enzyme must transcribe in the presence of structural barriers, such as nucleosomes, that can limit transcriptional progress and RNA throughput. We have initiated a major effort in my laboratory to understand how the eukaryotic polymerase negotiates transcription through these hurdles.
In principle, efficient transcription through nucleosomal barriers requires either template remodeling or an increase in the mechanical ability of the polymerase to overcome such barriers. We are investigating the transcription dynamics of single RNAP II molecules against force in the presence and absence of TFIIS, a transcription elongation factor known to increase transcription through nucleosomes. Using a single-molecule, dual-trap optical-tweezers assay combined with a novel method to enrich for active complexes, we found that the response of RNAP II to force is entirely determined by enzyme backtracking. Surprisingly, RNAP II molecules ceased to transcribe and were unable to recover from backtracks at a force of 7.5 ± 2 pN, only one-third of the force determined for Escherichia coli RNAP. We found that backtrack pause durations follow a t–3/2 power law, implying that during backtracking RNAP II diffuses in discrete base-pair steps and suggesting that backtracks may account for most of RNAP II pauses. Significantly, addition of TFIIS rescued backtracked enzymes and allowed transcription to proceed up to a force of 16.9 ± 3.4 piconewtons (pN). Together, these results describe a regulatory mechanism of transcription elongation in eukaryotes by which transcription factors modify the mechanical performance of RNAP II, allowing it to operate against higher loads and possible roadblocks, such as nucleosomes. We are now investigating transcription of nucleosomal DNA.
Mechanochemical Analysis of DNA Gyrase
Negative DNA supercoiling is essential in vivo to compact the genome, relieve torsional strain during replication, and promote local melting for vital processes such as transcript initiation by RNA polymerase. In bacteria, negative supercoiling is achieved through the activity of DNA gyrase, which uses the energy of ATP to work against mechanical stresses to drive the genome into an elastically strained configuration. Gyrase activity must compete kinetically with progression of the replication fork (which introduces positive supercoils) and the activity of topoisomerase I (which relaxes negative supercoils to help determine the steady-state level of supercoiling in the cell). The mechanical and kinetic properties of the gyrase motor are thus central to its in vivo role. Single-molecule techniques are uniquely suited to characterize these motor properties and have yielded important insights into the mechanisms of other topoisomerases, but had not been applied to DNA gyrase. Gyrase and other type II topoisomerases carry out a complex series of conformational changes that result in the passage of an intact DNA duplex (the T segment) through a transient break in another DNA duplex (the G segment), adding or removing two DNA links with each enzymatic cycle. The directionality of supercoiling is ensured by chiral wrapping of the DNA around a specialized domain of the enzyme prior to strand passage.
We have developed a single-molecule assay to investigate the mechanochemical cycle of gyrase. In this assay, we observe the activity of gyrase in real time by tracking the rotation of a submicron bead attached to the side of a stretched DNA molecule. In the presence of gyrase and ATP, bursts of rotation correspond to the processive, stepwise introduction of negative supercoils in strict multiples of two. We use tension to perturb and analyze the role of mechanical steps in the gyrase cycle. Changes in DNA tension have no detectable effect on the velocity of burst supercoiling, but the enzyme becomes markedly less processive as tension is increased over a range of only a few tenths of piconewtons. This behavior can be quantitatively explained by a simple mechanochemical model in which processivity depends on a kinetic competition between dissociation and rapid, tension-sensitive DNA wrapping. Moreover, using a high-resolution variant of our assay, we have detected pauses corresponding to slow kinetic steps within individual enzymatic cycles. The angular position of the dominant pause indicates that the rate-limiting step lies at the end of the reaction coordinate, subsequent to strand passage.
The RNA Translocation and Unwinding Mechanism of HCV NS3 Helicase and Its Coordination by ATP
Helicases are a ubiquitous class of enzymes involved in nearly all aspects of DNA and RNA metabolism. Despite recent progress in understanding their mechanism of action, limited resolution has left inaccessible the detailed mechanisms by which these enzymes couple the rearrangement of nucleic acid structures to ATP binding and hydrolysis. Observing individual mechanistic cycles of these motor proteins is central to understanding their cellular functions. We have been interested in understanding the molecular mechanisms underlying the activity of the hepatitis C virus (HCV) helicase (NS3) monomer. NS3, a key component of the HCV RNA replication machinery, lies in a membrane-bound complex with other proteins. It is a superfamily 2 (SF2) helicase essential for viral replication and therefore a potentially important drug target. NS3 possesses ATPase and 3' to 5' helicase activities, and it has been structurally characterized in various contexts.
We have developed a single-molecule assay to directly follow full-length NS3 moving on its RNA substrate. We use optical tweezers to apply a constant tension between two beads attached to the ends of a 60-bp RNA hairpin (Figure 1A) and monitor the RNA's end-to-end distance change as it is unwound by NS3. To interpret the enzyme's activity, we initially characterize the mechanical unfolding of the substrate in the absence of enzyme. The substrate unfolds at a force of 20.4 ± 0.2 pN (Figure 1B). When the instrument's force-feedback mechanism is used to hold the substrate at a constant force below 19 pN, no unfolding takes place over periods of several minutes. Therefore, substrate unfolding at external forces below 19 pN must be catalyzed by helicase. Using this assay, we have followed in real time, with 2-bp and 20-ms resolution, the RNA translocation and unwinding cycles of an HCV NS3 monomer. We show that the cyclic movement of NS3 is coordinated by ATP in discrete steps of 11 ± 3 bp and that unwinding occurs in rapid, smaller substeps of 3.6 ± 1.3 bp, also triggered by ATP binding, suggesting that NS3 moves like an inchworm. This ATP-coupling mechanism is likely to be applicable to other nonhexameric helicases involved in many essential cellular functions. Future studies of this enzyme will be directed to understanding the effect of RNA sequence and structure on its mechanism of translocation.
These projects have been supported in part by the National Institutes of Health and the Department of Energy.
Last updated: August 1, 2007
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