Hao Yu

The essential forces stabilizing membrane proteins and governing their folding and unfolding are difficult to decipher. Single-molecule atomic force spectroscopy mechanically unfolds individual membrane proteins and quantifies their dynamics and energetics. However, it remains challenging to structurally assign unfolding intermediates precisely and to deduce dominant interactions between specific residues that facilitate either the localized stabilization of these intermediates or the global assembly of membrane proteins. Here, we performed force spectroscopy experiments and multiscale molecular dynamics simulations to study the unfolding pathway of diacylglycerol kinase (DGK), a small trimeric multispan transmembrane enzyme. The remarkable agreement between experiments and simulations allowed precise structural assignment and interaction analysis of unfolding intermediates, bypassing existing limitations on structural mapping, and thus provided mechanistic explanations for the formation of these states. DGK unfolding was found to proceed with structural segments varying in size that do not correlate with its secondary structure. We identified intermolecular side-chain packing interactions as one of the major contributions to the stability of unfolding intermediates. Mutagenesis creating packing defects induced a dramatic decrease in the mechano-stability of corresponding intermediates and also in the thermo-stability of DGK trimer, in good agreement with predictions from simulations. Hence, the molecular determinants of the mechano- and thermo-stability of a membrane protein can be identified at residue resolution. The accurate structural assignment established and microscopic mechanism revealed in this work may substantially expand the scope of single-molecule studies of membrane proteins.

The forces that stabilize membrane proteins remain elusive to precise quantification. Particularly important, but poorly resolved, are the forces present during the initial unfolding of a membrane protein, where the most native set of interactions is present. A high‐precision, atomic force microscopy assay was developed to study the initial unfolding of bacteriorhodopsin. A rapid near‐equilibrium folding between the first three unfolding states was discovered, the two transitions corresponded to the unfolding of five and three amino acids, respectively, when using a cantilever optimized for 2 μs resolution. The third of these states was retinal‐stabilized and previously undetected, despite being the most mechanically stable state in the whole unfolding pathway, supporting 150 pN for more than 1 min. This ability to measure the dynamics of the initial unfolding of bacteriorhodopsin provides a platform for quantifying the energetics of membrane proteins under native‐like conditions.

Precisely quantifying the energetics that drive the folding of membrane proteins into a lipid bilayer remains challenging. More than 15 years ago, atomic force microscopy (AFM) emerged as a powerful tool to mechanically extract individual membrane proteins from a lipid bilayer. Concurrently, fluctuation theorems, such as the Jarzynski equality, were applied to deduce equilibrium free energies (ΔG₀) from non-equilibrium single-molecule force spectroscopy records. The combination of these two advances in single-molecule studies deduced the free-energy of the model membrane protein bacteriorhodopsin in its native lipid bilayer. To elucidate this free-energy landscape at a higher resolution, we applied two recent developments. First, as an input to the reconstruction, we used force-extension curves acquired with a 100-fold higher time resolution and 10-fold higher force precision than traditional AFM studies of membrane proteins. Next, by using an inverse Weierstrass transform and the Jarzynski equality, we removed the free energy associated with the force probe and determined the molecular free-energy landscape of the molecule under study, bacteriorhodopsin. The resulting landscape yielded an average unfolding free energy per amino acid (aa) of 1.0 ± 0.1 kcal/mol, in agreement with past single-molecule studies. Moreover, on a smaller spatial scale, this high-resolution landscape also agreed with an equilibrium measurement of a particular three-aa transition in bacteriorhodopsin that yielded 2.7 kcal/mol/aa, an unexpectedly high value. Hence, while average unfolding ΔG₀ per aa is a useful metric, the derived high-resolution landscape details significant local variation from the mean. More generally, we demonstrated that, as anticipated, the inverse Weierstrass transform is an efficient means to reconstruct free-energy landscapes from AFM data.

Pulling apart protein unfolding: Elucidating the details of how complex proteins fold is a longstanding challenge. Key insights into the unfolding pathways of diverse proteins have come from single-molecule force spectroscopy (SMFS) experiments in which proteins are literally pulled apart. Yu et al. developed a SMFS technique that could unfold individual bacteriorhodopsin molecules in a native lipid bilayer with 1-µs temporal resolution (see the Perspective by Müller and Gaub). The technique delivered a 100-fold improvement over earlier studies of bacteriorhodopsin and revealed many intermediates not seen before. The authors also observed unfolding and refolding transitions between intermediate states. Science , this issue p. 945 ; see also p. 907

How biomolecules fold: In order to fold, biomolecules must search a conformational energy landscape to find low-energy states. There are peaks in the landscape where the molecules must occupy unstable conformations for a short time. Neupane et al. used optical tweezers to observe these transition paths directly for single nucleic acid and protein molecules (see the Perspective by Wolynes). They measured a distribution of times taken to cross the transition path and found that the shape of the distribution agrees well with theory that assumes one-dimensional diffusion over the landscape. Science , this issue p. 239 ; see also p. 150

Significance: Structural transitions in proteins are characterized by the coefficient for intrachain diffusion, D , which determines the transition kinetics and reveals microscopic properties of the interactions governing folding. D has been measured for unfolded proteins and for native folding, but never for misfolding and aggregation, despite the importance of kinetics for driving these processes. We used single-molecule force spectroscopy to observe the misfolding of individual prion protein (PrP) molecules into stable, nonnative dimers. By reconstructing the energy landscape for dimer misfolding, we compared D for misfolding of PrP to that for native folding. Diffusion was 1,000-fold slower for misfolding, reflecting significant additional roughness in the energy landscape and confirming quantitatively the long-held hypothesis that misfolding landscapes are rougher than native landscapes.

The structural conversion of the prion protein PrP into a transmissible, misfolded form is the central element of prion disease, yet there is little consensus as to how it occurs. Key aspects of conversion into the diseased state remain unsettled, from details about the earliest stages of misfolding such as the involvement of partially- or fully-unfolded intermediates to the structure of the infectious state. Part of the difficulty in understanding the structural conversion arises from the complexity of the underlying energy landscapes. Single molecule methods provide a powerful tool for probing complex folding pathways as in prion misfolding, because they allow rare and transient events to be observed directly. We discuss recent work applying single-molecule probes to study misfolding in prion proteins, and what it has revealed about the folding dynamics of PrP that may underlie its unique behavior. We also discuss single-molecule studies probing the interactions that stabilize non-native structures within aggregates, pointing the way to future work that may help identify the microscopic events triggering pathogenic conversion. Although single-molecule approaches to misfolding are relatively young, they have a promising future in prion science.

Protein folding is described conceptually in terms of diffusion over a configurational free-energy landscape, typically reduced to a one-dimensional profile along a reaction coordinate. In principle, kinetic properties can be predicted directly from the landscape profile using Kramers theory for diffusive barrier crossing, including the folding rates and the transition time for crossing the barrier. Landscape theory has been widely applied to interpret the time scales for protein conformational dynamics, but protein folding rates and transition times have not been calculated directly from experimentally measured free-energy profiles. We characterized the energy landscape for native folding of the prion protein using force spectroscopy, measuring the change in extension of a single protein molecule at high resolution as it unfolded/refolded under tension. Key parameters describing the landscape profile were first recovered from the distributions of unfolding and refolding forces, allowing the diffusion constant for barrier crossing and the transition path time across the barrier to be calculated. The full landscape profile was then reconstructed from force-extension curves, revealing a double-well potential with an extended, partially unfolded transition state. The barrier height and position were consistent with the previous results. Finally, Kramers theory was used to predict the folding rates from the landscape profile, recovering the values observed experimentally both under tension and at zero force in ensemble experiments. These results demonstrate how advances in single-molecule theory and experiment are harnessing the power of landscape formalisms to describe quantitatively the mechanics of folding.

The duration of structural transitions in biopolymers is only a fraction of the time spent searching diffusively over the configurational energy landscape. We found the transition time, $\tau${}TP, and the diffusion constant, D, for DNA and RNA folding using energy landscapes obtained from single-molecule trajectories under tension in optical traps. DNA hairpins, RNA pseudoknots, and a riboswitch all had $\tau${}TP$\sim${}10\,\,$\mu${}s and D$\sim${}10-13\char21{}14\,\,m2/s, despite widely differing unfolding rates. These results show how energy-landscape analysis can be harnessed to characterize brief but critical events during folding reactions.