Single-molecule Force-clamp Spectroscopy Research Articles

Mechanochemistry: One Bond at a Time

Single-molecule force-clamp spectroscopy offers a novel platform for mechanically denaturing proteins by applying a constant force to a polyprotein. A powerful emerging application of the technique is that, by introducing a disulfide bond in each protein module, the chemical kinetics of disulfide bond cleavage under different stretching forces can be probed at the single-bond level. Even at forces much lower than that which can rupture the chemical bond, the breaking of the S-S bond at the presence of various chemical reducing agents is significantly accelerated. Our previous work demonstrated that the rate of thiol/disulfide exchange reaction is force-dependent and well-described by an Arrhenius term of the form r = A(exp((FΔx(r) - E(a))/k(B)T)[nucleophile]). From Arrhenius fits to the force dependency of the reduction rate, we measured the bond elongation parameter, Δx(r), along the reaction coordinate to the transition state of the S(N)2 reaction cleaved by different nucleophiles and enzymes, never before observed by any other technique. For S-S cleavage by various reducing agents, obtaining the Δx(r) value can help depicting the energy landscapes and elucidating the mechanisms of the reactions at the single-molecule level. Small nucleophiles, such as 1,4-dl-dithiothreitol (DTT), tris(2-carboxyethyl)phosphine (TCEP), and l-cysteine, react with the S-S bond with monotonically increasing rates under the applied force, while thioredoxin enzymes exhibit both stretching-favored and -resistant reaction-rate regimes. These measurements demonstrate the power of the single-molecule force-clamp spectroscopy approach in providing unprecedented access to chemical reactions.

Direct observation of an ensemble of stable collapsed states in the mechanical folding of ubiquitin.

Statistical theories of protein folding have long predicted plausible mechanisms for reducing the vast conformational space through distinct ensembles of structures. However, these predictions have remained untested by bulk techniques, because the conformational diversity of folding molecules has been experimentally unapproachable. Owing to recent advances in single molecule force-clamp spectroscopy, we are now able to probe the structure and dynamics of the small protein ubiquitin by measuring its length and mechanical stability during each stage of folding. Here, we discover that upon hydrophobic collapse, the protein rapidly selects a subset of minimum energy structures that are mechanically weak and essential precursors of the native fold. From this much reduced ensemble, the native state is acquired through a barrier-limited transition. Our results support the validity of statistical mechanics models in describing the folding of a small protein on biological timescales.

Osmolyte-induced separation of the mechanical folding phases of ubiquitin

Solvent molecules play key roles in the conformational dynamics of proteins. Here we use single molecule force-clamp spectroscopy to probe the role played by the stabilizing osmolyte glycerol on the conformational ensembles visited by a single ubiquitin protein folding after mechanical extension. Using a variety of force-pulse protocols, we find that glycerol stabilizes the native state of ubiquitin, making it more resistant to mechanical unfolding. We also find that although glycerol enhanced the hydrophobic collapse of unfolded and highly extended ubiquitins, it had no effect on the resulting collapsed states that are essential precursors of the folded state. These disparate effects of glycerol may be the result of distinct structural roles played by solvent molecules at the transition state of each folding ensemble. Our results open the way for a detailed analysis of the transition state structures that form along the folding trajectory of a mechanically extended protein.

Force-activated reactivity switch in a bimolecular chemical reaction

The effect of mechanical force on the free-energy surface that governs a chemical reaction is largely unknown. The combination of protein engineering with single-molecule force-clamp spectroscopy allows us to study the influence of mechanical force on the rate at which a protein disulfide bond is reduced by nucleophiles in a bimolecular substitution reaction (S(N)2). We found that cleavage of a protein disulfide bond by hydroxide anions exhibits an abrupt reactivity 'switch' at ∼500 pN, after which the accelerating effect of force on the rate of an S(N)2 chemical reaction greatly diminishes. We propose that an abrupt force-induced conformational change of the protein disulfide bond shifts its ground state, drastically changing its reactivity in S(N)2 chemical reactions. Our experiments directly demonstrate the action of a force-activated switch in the chemical reactivity of a single molecule.

Direct Identification of Two Distinct Transition State Structures in Reduction of a Disulfide Bond Revealed by Single Bond Force-clamp Spectroscopy

Disulfide bonds are common to many extracellular proteins, where they serve to stabilize the native conformation. Indeed, the thiol/disulfide exchange mechanism is involved in important and complex biological processes. Much experimental and theoretical work seems to support the idea that the reaction proceeds through an uncomplicated SN2. While in gas phase a double minimum potential model describes the potential energy surface governing the chemical reaction, in solution phase only unimodal, concerted profiles without intermediates have been identified. However, the detailed shapes of the energy surfaces of these reactions are largely unknown, because the collisions with solvent control the trajectories of the molecules. The combination of molecular engineering techniques with single molecule force-clamp spectroscopy has made it possible to monitor the reduction of single disulfide bonds, allowing us to experimentally measure the bond elongation at the reaction transition state with sub-Angstrom resolution. Such an experimental approach provides an unprecedented experimental platform to directly probe the energy landscape of a simple chemical reaction in solution at the single bond level. By greatly expanding the range of pulling forces up to 1.5 nN, where covalent bonds are not yet broken, here we demonstrate that the disulfide bond cleavage by hydroxyl occurs through a double-barrier energy landscape. Whereas at low pulling forces (100-500pN) the reaction rate is limited by a first energy barrier exhibiting a distance to the transition state Δx ∼0.5 A, at higher forces (500-1500 pN) a second energy barrier exhibiting a shorter transition state of Δx ∼0.1 A becomes dominant. Our experimental approach allows us to probe regions of the energy landscape that were previously experimentally inaccessible, revealing signatures of unanticipated complexity.

Revisiting Protein Folding at the Single Molecule Level

Determining the mechanism by which a protein folds remains a primary goal in biology. Statistical theories of protein folding have long predicted plausible mechanisms for reducing the vast conformational space through distinct ensembles of structures. However, these predictions have remained untested experimentally, since the multiplicity of trajectories and folding structures is averaged out using bulk techniques. Moreover, most intermediate conformations are only transiently present, rendering their isolation and characterization difficult by commonly used spectroscopic methods. Owing to recent advances in single molecule force-clamp spectroscopy, we are now able to probe the structure and dynamics of the small protein ubiquitin by measuring its length, mechanical stability and effect of solvent environment during each stage of folding. Here we discover that upon hydrophobic collapse, the protein rapidly selects a subset of non-native like, minimum energy structures that are mechanically weak and insensitive to the solvent environment. From this much reduced ensemble, the native state is acquired through a barrier-limited transition. The existence of such heterogeneous ensemble of minimum energy collapsed states was theoretically proposed by lattice simulations to be a milestone in the process of narrowing the available conformational space of a protein during its journey to the native fold, and a general feature of proteins that are naturally designed through evolution to fold on biological timescales. Here we demonstrate that such ensemble of collapsed states is also apparent in our experiments in the well-characterized I27 and Protein L proteins, albeit on different timescales, thus suggesting that their presence is ubiquitous to other mechanically stable proteins with a well-defined fold. Our results present the first experimental evidence for the validity of statistical mechanics models in describing the folding of small proteins on biological timescales.

Super-proteins From Fitness-threshold Selection Statistics

It is in principle possible to assign a fitness value (organism reproductive rate) to every sequence of any given protein. The resulting fitness landscape can be regarded as the convolution of two mappings: one from genotype to phenotype with another mapping from phenotype to fitness. Despite this intrinsic complexity, we may expect organism fitness to depend on protein properties bearing upon catalysis, interactions with other molecules, stability, et cetera. However, the specific relation between fitness, sequence space, evolution and protein molecular properties is not understood, a fact which hampers efforts to tap the enormous potential for protein and organism engineering contained in the exponentially-growing sequence databases. Here we show that a simple evolutionary hypothesis on the statistics of purifying natural selection over a fitness threshold is operational and leads to protein multi-feature optimization. We thus obtain variants of E. coli thioredoxin showing simultaneous, large-scale optimizing modulations in stability, folding/unfolding kinetics, bulk-solvent oxidoreductase activity and the two chemically and evolutionary different mechanisms of enzymatic catalysis revealed by single-molecule force clamp spectroscopy. Furthermore, preliminary experiments suggest that these variants may induce in-vivo resistance to thermal and oxidative stresses. We anticipate, therefore, applications in fields that involve organism engineering (microbial biotechonology, synthetic biology).

Solvent Bridging Determines The Molecular Architecture Of The Unfolding Transition State Of A Protein

Protecting osmolytes are ubiquitous in nature, where they play a vital role stabilizing intracellular proteins against adverse environmental conditions. While the solution thermodynamics of protein/osmolyte mixtures has been well characterized, information is lacking on how osmolytes influence the transition state structure and dynamics of proteins. Here we demonstrate a combination of single molecule force-clamp spectroscopy and solvent substitution that directly identifies the role of protecting osmolytes in the unfolding transition state structure of a protein. We measure the effect of osmolyte substitution on the rate of forced unfolding the I27 titin module. From the force dependency of the unfolding rate for each osmolyte we determine ΔGU, the height of the activation energy barrier, and ΔxU, the distance to the transition state. For all protecting osmolytes we measure ΔGU increases, demonstrating that the I27 protein is stabilized. More striking is the measurement of ΔxU. Unfolding the I27 protein in water gives a ΔxU = 2.5 A, a distance similar to the size of a water molecule. Water molecules have been identified as integral components of the unfolding transition state of the I27 protein, forming a solvent bridge between two β-strands. By varying osmolyte molecule size we rigorous test this solvent bridging hypothesis. We find that ΔxU correlates with osmolyte size for molecules ranging in size from 2.5 A to 5.6 A. However, for larger molecules (> 5.6 A) ΔxU remains unchanged relative to the value measured in water, suggesting these osmolytes do not participate in solvent bridging in the transition state. These studies uniquely probe the length scales over which solvent molecules can modify the molecular architecture of the unfolding transition of a protein, an area which remains beyond the reach of other experimental techniques.

Force-Clamp Spectroscopy Detects Residue Co-evolution in Enzyme Catalysis

Understanding how the catalytic mechanisms of enzymes are optimized through evolution remains a major challenge in molecular biology. The concept of co-evolution implicates that compensatory mutations occur to preserve the structure and function of proteins. We have combined statistical analysis of protein sequences with the sensitivity of single molecule force-clamp spectroscopy to probe how catalysis is affected by structurally distant correlated mutations in Escherichia coli thioredoxin. Our findings show that evolutionary anti-correlated mutations have an inhibitory effect on enzyme catalysis, whereas positively correlated mutations rescue the catalytic activity. We interpret these results in terms of an evolutionary tuning of both the enzyme-substrate binding process and the chemistry of the active site. Our results constitute a direct observation of distant residue co-evolution in enzyme catalysis.

Direct Observation of Markovian Behavior of the Mechanical Unfolding of Individual Proteins

Single-molecule force-clamp spectroscopy is a valuable tool to analyze unfolding kinetics of proteins. Previous force-clamp spectroscopy experiments have demonstrated that the mechanical unfolding of ubiquitin deviates from the generally assumed Markovian behavior and involves the features of glassy dynamics. Here we use single molecule force-clamp spectroscopy to study the unfolding kinetics of a computationally designed fast-folding mutant of the small protein GB1, which shares a similar β-grasp fold as ubiquitin. By treating the mechanical unfolding of polyproteins as the superposition of multiple identical Poisson processes, we developed a simple stochastic analysis approach to analyze the dwell time distribution of individual unfolding events in polyprotein unfolding trajectories. Our results unambiguously demonstrate that the mechanical unfolding of NuG2 fulfills all criteria of a memoryless Markovian process. This result, in contrast with the complex mechanical unfolding behaviors observed for ubiquitin, serves as a direct experimental demonstration of the Markovian behavior for the mechanical unfolding of a protein and reveals the complexity of the unfolding dynamics among structurally similar proteins. Furthermore, we extended our method into a robust and efficient pseudo-dwell-time analysis method, which allows one to make full use of all the unfolding events obtained in force-clamp experiments without categorizing the unfolding events. This method enabled us to measure the key parameters characterizing the mechanical unfolding energy landscape of NuG2 with improved precision. We anticipate that the methods demonstrated here will find broad applications in single-molecule force-clamp spectroscopy studies for a wide range of proteins.

Single-molecule avidin–biotin association reaction studied by force-clamp spectroscopy

Using single-molecule force-clamp spectroscopy, where the distance between the AFM tip and the sample surface is fixed and a few parallel avidin–biotin complexes are kept stretched by a certain force, we were able to observe the formation of single avidin–biotin bonds. Perspectives to use such an approach to study association reactions at single-molecule level in the conditions resembling those characteristic for some processes in vivo (e.g. virus-cell membrane attachment) are briefly discussed.

Dwell Time Analysis of a Single-Molecule Mechanochemical Reaction

Force-clamp spectroscopy is a novel technique for studying mechanochemistry at the single-bond level. Single disulfide bond reduction events are accurately detected as stepwise increases in the length of polyproteins that contain disulfide bonds and that are stretched at a constant force with the cantilever of an atomic force microscope (AFM). The kinetics of this reaction has been measured from single-exponential fits to ensemble averages of the reduction events. However, exponential fits are notoriously ambiguous to use in cases of kinetic data showing multiple reaction pathways. Here we introduce a dwell time analysis technique, of widespread use in the single ion channel field, that we apply to the examination of the kinetics of reduction of disulfide bonds measured from single-molecule force-clamp spectroscopy traces. In this technique, exponentially distributed dwell time data is plotted as a histogram with a logarithmic time scale and a square root ordinate. The advantage of logarithmic histograms is that exponentially distributed dwell times appear as well-defined peaks in the distribution, greatly enhancing our ability to detect multiple kinetic pathways. We apply this technique to examine the distribution of dwell times of 4488 single disulfide bond reduction events measured in the presence of two very different kinds of reducing agents: tris-(2-carboxyethyl)phosphine hydrochloride (TCEP) and the enzyme thioredoxin (TRX). A different clamping force is used for each reducing agent to obtain distributions of dwell times on a similar time scale. In the case of TCEP, the logarithmic histogram of dwell times showed a single peak, corresponding to a single reaction mechanism. By contrast, similar experiments done with TRX showed two well-separated peaks, marking two distinct modes of chemical reduction operating simultaneously. These experiments demonstrate that dwell time analysis techniques are a powerful approach to studying chemical reactions at the single-molecule level.

Probing the chemistry of thioredoxin catalysis with force

Thioredoxins are enzymes that catalyse disulphide bond reduction in all living organisms. Although catalysis is thought to proceed through a substitution nucleophilic bimolecular (S(N)2) reaction, the role of the enzyme in modulating this chemical reaction is unknown. Here, using single-molecule force-clamp spectroscopy, 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 A (+/- s.e.m.), and the second elongating the substrate disulphide bond by 0.17 +/- 0.02 A (+/- 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. 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.

Force-dependent chemical kinetics of disulfide bond reduction observed with single-molecule techniques

The mechanism by which mechanical force regulates the kinetics of a chemical reaction is unknown. Here, we use single-molecule force-clamp spectroscopy and protein engineering to study the effect of force on the kinetics of thiol/disulfide exchange. Reduction of disulfide bonds through the thiol/disulfide exchange chemical reaction is crucial in regulating protein function and is known to occur in mechanically stressed proteins. We apply a constant stretching force to single engineered disulfide bonds and measure their rate of reduction by DTT. Although the reduction rate is linearly dependent on the concentration of DTT, it is exponentially dependent on the applied force, increasing 10-fold over a 300-pN range. This result predicts that the disulfide bond lengthens by 0.34 A at the transition state of the thiol/disulfide exchange reaction. Our work at the single bond level directly demonstrates that thiol/disulfide exchange in proteins is a force-dependent chemical reaction. Our findings suggest that mechanical force plays a role in disulfide reduction in vivo, a property that has never been explored by traditional biochemistry. Furthermore, our work also indicates that the kinetics of any chemical reaction that results in bond lengthening will be force-dependent.

Single-molecule force spectroscopy reveals signatures of glassy dynamics in the energy landscape of ubiquitin

The conformational energy landscape of a protein out of equilibrium is poorly understood. We use single-molecule force-clamp spectroscopy to measure the kinetics of unfolding of the protein ubiquitin under a constant force. We discover a surprisingly broad distribution of unfolding rates that follows a power law with no characteristic mean. The structural fluctuations that give rise to this distribution reveal the architecture of the protein’s energy landscape. Following models of glassy dynamics, this complex kinetics implies large fluctuations in the energies of the folded protein, characterized by an exponential distribution with a width of 5–10kBT. Our results predict the existence of a ‘glass transition’ force below which the folded conformations interconvert between local minima on multiple timescales. These techniques offer a new tool to further test statistical energy landscape theories experimentally.