Thioredoxins are enzymes that catalyse disulphide bond reduction in all living organisms 1 . Although catalysis is thought to proceed through a substitution nucleophilic bimolecular (S N 2) reaction 1,2 , the role of the enzyme in modulating this chemical reaction is unknown. Here, using single-molecule force-clamp spectroscopy 3,4 , we investigate the catalytic mechanism of Escherichia coli thioredoxin (Trx). We applied mechanical force in the range of 25-600 pN to a disulphide bond substrate and monitored the reduction of these bonds by individual enzymes. We detected two alternative forms of the catalytic reaction, the first requiring a reorientation of the substrate disulphide bond, causing a shortening of the substrate polypeptide by 0.79 ± 0.09 Å (± s.e.m.), and the second elongating the substrate disulphide bond by 0.17 ± 0.02 Å (±s.e.m.). These results support the view that the Trx active site regulates the geometry of the participating sulphur atoms with sub-ångström precision to achieve efficient catalysis. Our results indicate that substrate conformational changes may be important in the regulation of Trx activity under conditions of oxidative stress and mechanical injury, such as those experienced in cardiovascular disease 5,6 . Furthermore, single-molecule atomic force microscopy techniques, as shown here, can probe dynamic rearrangements within an enzyme's active site during catalysis that cannot be resolved with any other current structural biological technique.One of the principal challenges of understanding enzyme catalysis, a central problem in biology, is resolving the dynamics of enzyme-substrate interactions with sub-ångström resolution-the length scale at which chemistry occurs 7 . Although nuclear magnetic resonance (NMR) and X-ray crystallography determinations of protein structures can reach down to the sub-ångström level, they cannot yet provide dynamic information about enzyme

The introduction of disulfide bonds into proteins creates additional mechanical barriers and limits the unfolded contour length (i.e., the maximal extension) measured by single-molecule force spectroscopy. Here, we engineer single disulfide bonds into four different locations of the human cardiac titin module (I27) to control the contour length while keeping the distance to the transition state unchanged. This enables the study of several biologically important parameters. First, we are able to precisely determine the end-to-end length of the transition state before unfolding (53 Angstrom), which is longer than the end-to-end length of the protein obtained from NMR spectroscopy (43 Angstrom). Second, the measured contour length per amino acid from five different methods (4.0 +/- 0.2 Angstrom) is longer than the end-to-end length obtained from the crystal structure (3.6 Angstrom). Our measurement of the contour length takes into account all the internal degrees of freedom of the polypeptide chain, whereas crystallography measures the end-to-end length within the "frozen" protein structure. Furthermore, the control of contour length and therefore the number of amino acids unraveled before reaching the disulfide bond (n) facilitates the test of the chain length dependence on the folding time (tau(F)). We find that both a power law scaling tau(F) lambda n(lambda) with lambda = 4.4, and an exponential scaling with n(0.6) fit the data range, in support of different protein-folding scenarios.

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We unfold and extend single proteins at a high force and then linearly relax the force to probe their collapse mechanisms. We observe a large variability in the extent of their recoil. Although chain entropy makes a small contribution, we show that the observed variability results from hydrophobic interactions with randomly varying magnitude from protein to protein. This collapse mechanism is common to highly extended proteins, including nonfolding elastomeric proteins like PEVK from titin. Our observations explain the puzzling differences between the folding behavior of highly extended proteins, from those folding after chemical or thermal denaturation. Probing the collapse of highly extended proteins with force spectroscopy allows separation of the different driving forces in protein folding.atomic force microscopy ͉ molecular dynamics ͉ protein folding ͉ single molecule P roteins can reversibly fold from a random coil conformation into a well defined native state, a process constituting a major research area in biology. Traditionally, experiments involved varying the ambient environment, such as changing the temperature or pressure, or using denaturing chemicals. Protein folding probed under these conditions has revealed two-state folding for many small proteins (1-3). Based on such experiments, Wolynes and colleagues proposed that the energy landscape of a collapsing polypeptide is funnel-shaped under folding conditions (4-6). In this scenario, the protein's energy decreases as it forms favorable interactions, thus driving it toward the native state. In the classic folding experiments, as well as in the theoretical models, proteins start in the denatured state from collapsed random coil conformations that are only a few Ångström larger than their native state (7,8). In these conformations, the side chains of the collapsed polypeptide are in close proximity to each other. It is widely accepted that, under such conditions, protein folding is driven mostly by hydrophobic interactions that are finely balanced by entropy (9-11). However, given that the denatured state in these experiments is not well defined, it has proved difficult to separate the hydrophobic, electrostatic, and entropic contributions to protein collapse and folding. We use single-molecule force-clamp spectroscopy to bring proteins to an extended conformation of Ͼ80% of their contour length, where the side chains are separated and exposed to the solvent, and native contact formation is rare (12-14). Thus, proteins are driven to the outer regions of the folding landscape, which have not been explored so far. Studying the collapse of such extended proteins greatly simplifies the folding dynamics, permitting a more direct identification of the major driving forces (11).Highly extended proteins have been routinely described as entropic chains using models of polymer elasticity such as the worm-like chain (WLC) model (15) or the freely rotating chain model (16). In this simplified picture, the collapse of a protein from an extended state is driven by...

Using the recently developed single molecule force-clamp technique we quantitatively measure the kinetics of conformational changes of polyprotein molecules at a constant force. In response to an applied force of 110 pN, we measure the dwell times of 1647 unfolding events of individual ubiquitin modules within each protein chain. We then establish a rigorous method for analyzing force-clamp data using order statistics. This allows us to test the success of a history-independent, two-state model in describing the kinetics of the unfolding process. We find that the average unfolding trajectory is independent of the number of protein modules N in each trajectory, which varies between 3 and 12 (the engineered protein length), suggesting that the unfolding events in each chain are uncorrelated. We then derive a binomial distribution of dwell times to describe the stochastic dynamics of protein unfolding. This distribution successfully describes 81% of the data with a single rate constant of alpha = 0.6 s(-1) for all N. The remainder of the data that cannot be accounted for suggests alternative unfolding barriers in the energy landscape of the protein. This method investigates the statistical features of unfolding beyond the average measurement of a single rate constant, thus providing an attractive alternative for measuring kinetics by force-clamp spectroscopy.