Quantifying a light-induced energetic change in bacteriorhodopsin by force spectroscopy.

Ligand-induced conformational changes are critical to the function of many membrane proteins and arise from numerous intramolecular interactions. In the photocycle of the model membrane protein bacteriorhodopsin (bR), absorption of a photon by retinal triggers a conformational cascade that results in pumping a proton across the cell membrane. While decades of spectroscopy and structural studies have probed this photocycle in intricate detail, changes in intramolecular energetics that underlie protein motions have remained elusive to experimental quantification. Here, we measured these energetics on the millisecond time scale using atomic-force-microscopy-based single-molecule force spectroscopy. Precisely, timed light pulses triggered the bR photocycle while we measured the equilibrium unfolding and refolding of the terminal 8-amino-acid region of bR's G-helix. These dynamics changed when the EF-helix pair moved ~9 Å away from this end of the G helix during the "open" portion of bR's photocycle. In ~60% of the data, we observed abrupt light-induced destabilization of 3.4 ± 0.3 kcal/mol, lasting 38 ± 3 ms. The kinetics and pH-dependence of this destabilization were consistent with prior measurements of bR's open phase. The frequency of light-induced destabilization increased with the duration of illumination and was dramatically reduced in the triple mutant (D96G/F171C/F219L) thought to trap bR in its open phase. In the other ~40% of the data, photoexcitation unexpectedly stabilized a longer-lived putative misfolded state. Through this work, we establish a general single-molecule force spectroscopy approach for measuring ligand-induced energetics and lifetimes in membrane proteins.

Force-Activated DNA Substrates for In Situ Generation of ssDNA and Designed ssDNA/dsDNA Structures in an Optical-Trapping Assay.

Single-molecule force spectroscopy can precisely probe the biomechanical interactions of proteins that unwind duplex DNA and bind to and wrap around single-stranded (ss)DNA. Yet assembly of the required substrates, which often contain a ssDNA segment embedded within a larger double-stranded (ds)DNA construct, can be time-consuming and inefficient, particularly when using a standard three-way hybridization protocol. In this chapter, we detail how to construct a variety of force-activated DNA substrates more efficiently. To do so, we engineered a dsDNA molecule with a designed sequence of specified GC content positioned between two enzymatically induced, site-specific nicks. Partially pulling this substrate into the overstretching transition of DNA (~65 pN) using an optical trap led to controlled dissociation of the ssDNA segment delineated by the two nicks. Here, we describe protocols for generating ssDNA of up to 1000 nucleotides as well as more complex structures, such as a 120-base-pair DNA hairpin positioned next to a 33-nucleotide ssDNA segment. The utility of the hairpin substrate was demonstrated by measuring the motion of E. coli. RecQ, a 3'-to-5' DNA helicase.

Free-energy changes of bacteriorhodopsin point mutants measured by single-molecule force spectroscopy.

Single amino acid mutations provide quantitative insight into the energetics that underlie the dynamics and folding of membrane proteins. Chemical denaturation is the most widely used assay and yields the change in unfolding free energy (ΔΔG). It has been applied to >80 different residues of bacteriorhodopsin (bR), a model membrane protein. However, such experiments have several key limitations: 1) a nonnative lipid environment, 2) a denatured state with significant secondary structure, 3) error introduced by extrapolation to zero denaturant, and 4) the requirement of globally reversible refolding. We overcame these limitations by reversibly unfolding local regions of an individual protein with mechanical force using an atomic-force-microscope assay optimized for 2 µs time resolution and 1 pN force stability. In this assay, bR was unfolded from its native bilayer into a well-defined, stretched state. To measure ΔΔG, we introduced two alanine point mutations into an 8-amino-acid region at the C-terminal end of bR's G helix. For each, we reversibly unfolded and refolded this region hundreds of times while the rest of the protein remained folded. Our single-molecule-derived ΔΔG for mutant L223A (-2.3 ± 0.6 kcal/mol) quantitatively agreed with past chemical denaturation results while our ΔΔG for mutant V217A was 2.2-fold larger (-2.4 ± 0.6 kcal/mol). We attribute the latter result, in part, to contact between Val217 and a natively bound squalene lipid, highlighting the contribution of membrane protein-lipid contacts not present in chemical denaturation assays. More generally, we established a platform for determining ΔΔG for a fully folded membrane protein embedded in its native bilayer.

Modulation of a protein-folding landscape revealed by AFM-based force spectroscopy notwithstanding instrumental limitations.

Single-molecule force spectroscopy is a powerful tool for studying protein folding. Over the last decade, a key question has emerged: how are changes in intrinsic biomolecular dynamics altered by attachment to µm-scale force probes via flexible linkers? Here, we studied the folding/unfolding of α3D using atomic force microscopy (AFM)-based force spectroscopy. α3D offers an unusual opportunity as a prior single-molecule fluorescence resonance energy transfer (smFRET) study showed α3D's configurational diffusion constant within the context of Kramers theory varies with pH. The resulting pH dependence provides a test for AFM-based force spectroscopy's ability to track intrinsic changes in protein folding dynamics. Experimentally, however, α3D is challenging. It unfolds at low force (<15 pN) and exhibits fast-folding kinetics. We therefore used focused ion beam-modified cantilevers that combine exceptional force precision, stability, and temporal resolution to detect state occupancies as brief as 1 ms. Notably, equilibrium and nonequilibrium force spectroscopy data recapitulated the pH dependence measured using smFRET, despite differences in destabilization mechanism. We reconstructed a one-dimensional free-energy landscape from dynamic data via an inverse Weierstrass transform. At both neutral and low pH, the resulting constant-force landscapes showed minimal differences (∼0.2 to 0.5 kBT) in transition state height. These landscapes were essentially equal to the predicted entropic barrier and symmetric. In contrast, force-dependent rates showed that the distance to the unfolding transition state increased as pH decreased and thereby contributed to the accelerated kinetics at low pH. More broadly, this precise characterization of a fast-folding, mechanically labile protein enables future AFM-based studies of subtle transitions in mechanoresponsive proteins.

Type III secretion system effector proteins are mechanically labile.

Multiple gram-negative bacteria encode type III secretion systems (T3SS) that allow them to inject effector proteins directly into host cells to facilitate colonization. To be secreted, effector proteins must be at least partially unfolded to pass through the narrow needle-like channel (diameter <2 nm) of the T3SS. Fusion of effector proteins to tightly packed proteins-such as GFP, ubiquitin, or dihydrofolate reductase (DHFR)-impairs secretion and results in obstruction of the T3SS. Prior observation that unfolding can become rate-limiting for secretion has led to the model that T3SS effector proteins have low thermodynamic stability, facilitating their secretion. Here, we first show that the unfolding free energy ([Formula: see text]) of two Salmonella effector proteins, SptP and SopE2, are 6.9 and 6.0 kcal/mol, respectively, typical for globular proteins and similar to published [Formula: see text] for GFP, ubiquitin, and DHFR. Next, we mechanically unfolded individual SptP and SopE2 molecules by atomic force microscopy (AFM)-based force spectroscopy. SptP and SopE2 unfolded at low force (Funfold ≤ 17 pN at 100 nm/s), making them among the most mechanically labile proteins studied to date by AFM. Moreover, their mechanical compliance is large, as measured by the distance to the transition state (Δx‡ = 1.6 and 1.5 nm for SptP and SopE2, respectively). In contrast, prior measurements of GFP, ubiquitin, and DHFR show them to be mechanically robust (Funfold > 80 pN) and brittle (Δx‡ < 0.4 nm). These results suggest that effector protein unfolding by T3SS is a mechanical process and that mechanical lability facilitates efficient effector protein secretion.

Correcting molecular transition rates measured by single-molecule force spectroscopy for limited temporal resolution.

Equilibrium free-energy-landscape parameters governing biomolecular folding can be determined from nonequilibrium force-induced unfolding by measuring the rates k for transitioning back and forth between states as a function of force F. However, bias in the observed forward and reverse rates is introduced by limited effective temporal resolution, which includes the mechanical response time of the force probe and any smoothing used to improve the signal-to-noise ratio. Here we use simulations to characterize this bias, which is most prevalent when the ratio of forward and reverse rates is far from unity. We find deviations in k(F) at high rates, due to unobserved transitions from short- to long-lived states, and at low rates, due to the corresponding unobserved transitions from long- to short-lived states. These missing events introduce erroneous curvature in log(k) vs F that leads to incorrect landscape parameter determination. To correct the measured k(F), we derive a pair of model-independent analytical formulas. The first correction accounts for unobserved transitions from short- to long-lived states, but does surprisingly little to correct the erroneous energy-landscape parameters. Only by subsequently applying the second formula, which corrects the corresponding reverse process, do we recover the expected k(F) and energy-landscape quantities. Going forward, these corrections should be applied to transition-rate data whenever the highest measured rate is not at least an order of magnitude slower than the effective temporal resolution.

Quantifying the Native Energetics Stabilizing Bacteriorhodopsin by Single-Molecule Force Spectroscopy.

We quantified the equilibrium (un)folding free energy ΔG_{0} of an eight-amino-acid region starting from the fully folded state of the model membrane-protein bacteriorhodopsin using single-molecule force spectroscopy. Analysis of equilibrium and nonequilibrium data yielded consistent, high-precision determinations of ΔG_{0} via multiple techniques (force-dependent kinetics, Crooks fluctuation theorem, and inverse Boltzmann analysis). We also deduced the full 1D projection of the free-energy landscape in this region. Importantly, ΔG_{0} was determined in bacteriorhodopsin's native bilayer, an advance over traditional results obtained by chemical denaturation in nonphysiological detergent micelles.

Bending and looping of long DNA by Polycomb repressive complex 2 revealed by AFM imaging in liquid.

Polycomb repressive complex 2 (PRC2) is a histone methyltransferase that methylates histone H3 at Lysine 27. PRC2 is critical for epigenetic gene silencing, cellular differentiation and the formation of facultative heterochromatin. It can also promote or inhibit oncogenesis. Despite this importance, the molecular mechanisms by which PRC2 compacts chromatin are relatively understudied. Here, we visualized the binding of PRC2 to naked DNA in liquid at the single-molecule level using atomic force microscopy. Analysis of the resulting images showed PRC2, consisting of five subunits (EZH2, EED, SUZ12, AEBP2 and RBBP4), bound to a 2.5-kb DNA with an apparent dissociation constant ($K_{\rm{D}}^{{\rm{app}}}$) of 150 ± 12 nM. PRC2 did not show sequence-specific binding to a region of high GC content (76%) derived from a CpG island embedded in such a long DNA substrate. At higher concentrations, PRC2 compacted DNA by forming DNA loops typically anchored by two or more PRC2 molecules. Additionally, PRC2 binding led to a 3-fold increase in the local bending of DNA's helical backbone without evidence of DNA wrapping around the protein. We suggest that the bending and looping of DNA by PRC2, independent of PRC2's methylation activity, may contribute to heterochromatin formation and therefore epigenetic gene silencing.

Membrane-Protein Unfolding Intermediates Detected with Enhanced Precision Using a Zigzag Force Ramp.

Precise quantification of the energetics and interactions that stabilize membrane proteins in a lipid bilayer is a long-sought goal. Toward this end, atomic force microscopy has been used to unfold individual membrane proteins embedded in their native lipid bilayer, typically by retracting the cantilever at a constant velocity. Recently, unfolding intermediates separated by as few as two amino acids were detected using focused-ion-beam-modified ultrashort cantilevers. However, unambiguously discriminating between such closely spaced states remains challenging, in part because any individual unfolding trajectory only occupies a subset of the total number of intermediates. Moreover, structural assignment of these intermediates via worm-like-chain analysis is hindered by brief dwell times compounded with thermal and instrumental noise. To overcome these issues, we moved the cantilever in a sawtooth pattern of 6-12 nm, offset by 0.25-1 nm per cycle, generating a "zigzag" force ramp of alternating positive and negative loading rates. We applied this protocol to the model membrane protein bacteriorhodopsin (bR). In contrast to conventional studies that extract bR's photoactive retinal along with the first transmembrane helix, we unfolded bR in the presence of its retinal. To do so, we introduced a previously developed enzymatic-cleavage site between helices E and F and pulled from the top of the E helix using a site-specific, covalent attachment. The resulting zigzag unfolding trajectories occupied 40% more states per trajectory and occupied those states for longer times than traditional constant-velocity records. In total, we identified 31 intermediates during the unfolding of five helices of EF-cleaved bR. These included a previously reported, mechanically robust intermediate located between helices C and B that, with our enhanced resolution, is now shown to be two distinct states separated by three amino acids. Interestingly, another intermediate directly interacted with the retinal, an interaction confirmed by removing the retinal.

Imaging DNA Equilibrated onto Mica in Liquid Using Biochemically Relevant Deposition Conditions.

For over 25 years, imaging of DNA by atomic force microscopy has been intensely pursued. Ideally, such images are then used to probe the physical properties of DNA and characterize protein-DNA interactions. The atomic flatness of mica makes it the preferred substrate for high signal-to-noise ratio (SNR) imaging, but the negative charge of mica and DNA hinders deposition. Traditional methods for imaging DNA and protein-DNA complexes in liquid have drawbacks: DNA conformations with an anomalous persistence length ( p), low SNR, and/or ionic deposition conditions detrimental to preserving protein-DNA interactions. Here, we developed a process to bind DNA to mica in a buffer containing both MgCl2 and KCl that resulted in high SNR images of equilibrated DNA in liquid. Achieving an equilibrated 2D configuration ( i. e., p = 50 nm) not only implied a minimally perturbative binding process but also improved data quality and quantity because the DNA's configuration was more extended. In comparison to a purely NiCl2-based protocol, we showed that an 8-fold larger fraction (90%) of 680-nm-long DNA molecules could be quantified. High-resolution images of select equilibrated molecules revealed the right-handed structure of DNA with a helical pitch of 3.5 nm. Deposition and imaging of DNA was achieved over a wide range of monovalent and divalent ionic conditions, including a buffer containing 50 mM KCl and 3 mM MgCl2. Finally, we imaged two protein-DNA complexes using this protocol: a restriction enzyme bound to DNA and a small three-nucleosome array. We expect such deposition of protein-DNA complexes at biochemically relevant ionic conditions will facilitate biophysical insights derived from imaging diverse protein-DNA complexes.

Quantifying the Initial Unfolding of Bacteriorhodopsin Reveals Retinal Stabilization.

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.

High-Precision Single-Molecule Characterization of the Folding of an HIV RNA Hairpin by Atomic Force Microscopy.

The folding of RNA into a wide range of structures is essential for its diverse biological functions from enzymatic catalysis to ligand binding and gene regulation. The unfolding and refolding of individual RNA molecules can be probed by single-molecule force spectroscopy (SMFS), enabling detailed characterization of the conformational dynamics of the molecule as well as the free-energy landscape underlying folding. Historically, high-precision SMFS studies of RNA have been limited to custom-built optical traps. Although commercial atomic force microscopes (AFMs) are widely deployed and offer significant advantages in ease-of-use over custom-built optical traps, traditional AFM-based SMFS lacks the sensitivity and stability to characterize individual RNA molecules precisely. Here, we developed a high-precision SMFS assay to study RNA folding using a commercial AFM and applied it to characterize a small RNA hairpin from HIV that plays a key role in stimulating programmed ribosomal frameshifting. We achieved rapid data acquisition in a dynamic assay, unfolding and then refolding the same individual hairpin more than 1,100 times in 15 min. In comparison to measurements using optical traps, our AFM-based assay featured a stiffer force probe and a less compliant construct, providing a complementary measurement regime that dramatically accelerated equilibrium folding dynamics. Not only did kinetic analysis of equilibrium trajectories of the HIV RNA hairpin yield the traditional parameters used to characterize folding by SMFS (zero-force rate constants and distances to the transition state), but we also reconstructed the full 1D projection of the folding free-energy landscape comparable to state-of-the-art studies using dual-beam optical traps, a first for this RNA hairpin and AFM studies of nucleic acids in general. Looking forward, we anticipate that the ease-of-use of our high-precision assay implemented on a commercial AFM will accelerate studying folding of diverse nucleic acid structures.

FEATHER: Automated Analysis of Force Spectroscopy Unbinding and Unfolding Data via a Bayesian Algorithm.

Single-molecule force spectroscopy (SMFS) provides a powerful tool to explore the dynamics and energetics of individual proteins, protein-ligand interactions, and nucleic acid structures. In the canonical assay, a force probe is retracted at constant velocity to induce a mechanical unfolding/unbinding event. Next, two energy landscape parameters, the zero-force dissociation rate constant (ko) and the distance to the transition state (Δx‡), are deduced by analyzing the most probable rupture force as a function of the loading rate, the rate of change in force. Analyzing the shape of the rupture force distribution reveals additional biophysical information, such as the height of the energy barrier (ΔG‡). Accurately quantifying such distributions requires high-precision characterization of the unfolding events and significantly larger data sets. Yet, identifying events in SMFS data is often done in a manual or semiautomated manner and is obscured by the presence of noise. Here, we introduce, to our knowledge, a new algorithm, FEATHER (force extension analysis using a testable hypothesis for event recognition), to automatically identify the locations of unfolding/unbinding events in SMFS records and thereby deduce the corresponding rupture force and loading rate. FEATHER requires no knowledge of the system under study, does not bias data interpretation toward the dominant behavior of the data, and has two easy-to-interpret, user-defined parameters. Moreover, it is a linear algorithm, so it scales well for large data sets. When analyzing a data set from a polyprotein containing both mechanically labile and robust domains, FEATHER featured a 30-fold improvement in event location precision, an eightfold improvement in a measure of the accuracy of the loading rate and rupture force distributions, and a threefold reduction of false positives in comparison to two representative reference algorithms. We anticipate FEATHER being leveraged in more complex analysis schemes, such as the segmentation of complex force-extension curves for fitting to worm-like chain models and extended in future work to data sets containing both unfolding and refolding transitions.

Improved free-energy landscape reconstruction of bacteriorhodopsin highlights local variations in unfolding energy.

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.

Improved Free-Energy Landscape Quantification Illustrated with a Computationally Designed Protein-Ligand Interaction.

Quantifying the energy landscape underlying protein-ligand interactions leads to an enhanced understanding of molecular recognition. A powerful yet accessible single-molecule technique is atomic force microscopy (AFM)-based force spectroscopy, which generally yields the zero-force dissociation rate constant (koff ) and the distance to the transition state (Δx≠ ). Here, we introduce an enhanced AFM assay and apply it to probe the computationally designed protein DIG10.3 binding to its target ligand, digoxigenin. Enhanced data quality enabled an analysis that yielded the height of the transition state (ΔG≠ =6.3±0.2 kcal mol-1 ) and the shape of the energy barrier at the transition state (linear-cubic) in addition to the traditional parameters [koff (=4±0.1×10-4  s-1 ) and Δx≠ (=8.3±0.1 Å)]. We expect this automated and relatively rapid assay to provide a more complete energy landscape description of protein-ligand interactions and, more broadly, the diverse systems studied by AFM-based force spectroscopy.

Going Vertical To Improve the Accuracy of Atomic Force Microscopy Based Single-Molecule Force Spectroscopy.

Single-molecule force spectroscopy (SMFS) is a powerful technique to characterize the energy landscape of individual proteins, the mechanical properties of nucleic acids, and the strength of receptor-ligand interactions. Atomic force microscopy (AFM)-based SMFS benefits from ongoing progress in improving the precision and stability of cantilevers and the AFM itself. Underappreciated is that the accuracy of such AFM studies remains hindered by inadvertently stretching molecules at an angle while measuring only the vertical component of the force and extension, degrading both measurements. This inaccuracy is particularly problematic in AFM studies using double-stranded DNA and RNA due to their large persistence length (p ≈ 50 nm), often limiting such studies to other SMFS platforms (e.g., custom-built optical and magnetic tweezers). Here, we developed an automated algorithm that aligns the AFM tip above the DNA's attachment point to a coverslip. Importantly, this algorithm was performed at low force (10-20 pN) and relatively fast (15-25 s), preserving the connection between the tip and the target molecule. Our data revealed large uncorrected lateral offsets for 100 and 650 nm DNA molecules [24 ± 18 nm (mean ± standard deviation) and 180 ± 110 nm, respectively]. Correcting this offset yielded a 3-fold improvement in accuracy and precision when characterizing DNA's overstretching transition. We also demonstrated high throughput by acquiring 88 geometrically corrected force-extension curves of a single individual 100 nm DNA molecule in ∼40 min and versatility by aligning polyprotein- and PEG-based protein-ligand assays. Importantly, our software-based algorithm was implemented on a commercial AFM, so it can be broadly adopted. More generally, this work illustrates how to enhance AFM-based SMFS by developing more sophisticated data-acquisition protocols.

Force Spectroscopy with 9-µs Resolution and Sub-pN Stability by Tailoring AFM Cantilever Geometry.

Atomic force microscopy (AFM)-based single-molecule force spectroscopy (SMFS) is a powerful yet accessible means to characterize the unfolding/refolding dynamics of individual molecules and resolve closely spaced, transiently occupied folding intermediates. On a modern commercial AFM, these applications and others are now limited by the mechanical properties of the cantilever. Specifically, AFM-based SMFS data quality is degraded by a commercial cantilever's limited combination of temporal resolution, force precision, and force stability. Recently, we modified commercial cantilevers with a focused ion beam to optimize their properties for SMFS. Here, we extend this capability by modifying a 40 × 18 µm2 cantilever into one terminated with a gold-coated, 4 × 4 µm2 reflective region connected to an uncoated 2-µm-wide central shaft. This "Warhammer" geometry achieved 8.5-µs resolution coupled with improved force precision and sub-pN stability over 100 s when measured on a commercial AFM. We highlighted this cantilever's biological utility by first resolving a calmodulin unfolding intermediate previously undetected by AFM and then measuring the stabilization of calmodulin by myosin light chain kinase at dramatically higher unfolding velocities than in previous AFM studies. More generally, enhancing data quality via an improved combination of time resolution, force precision, and force stability will broadly benefit biological applications of AFM.

Force-activated DNA substrates for probing individual proteins interacting with single-stranded DNA.

Single-molecule force spectroscopy provides insight into how proteins bind to and move along DNA. Such studies often embed a single-stranded (ss) DNA region within a longer double-stranded (ds) DNA molecule. Yet, producing these substrates remains laborious and inefficient, particularly when using the traditional three-way hybridization. Here, we developed a force-activated substrate that yields an internal 1000 nucleotide (nt) ssDNA region when pulled partially into the overstretching transition (∼65 pN) by engineering a 50%-GC segment to have no adjacent GC base pairs. Once the template was made, these substrates were efficiently prepared by polymerase chain reaction amplification followed by site-specific nicking. We also generated a more complex structure used in high-resolution helicase studies, a DNA hairpin adjacent to 33 nt of ssDNA. The temporally defined generation of individual hairpin substrates in the presence of RecQ helicase and saturating adenine triphosphate let us deduce that RecQ binds to ssDNA via a near diffusion-limited reaction. More broadly, these substrates enable the precise initiation of an important class of protein-DNA interactions.

Rapid Characterization of a Mechanically Labile α-Helical Protein Enabled by Efficient Site-Specific Bioconjugation.

Atomic force microscopy (AFM)-based single-molecule force spectroscopy (SMFS) is a powerful yet accessible means to characterize mechanical properties of biomolecules. Historically, accessibility relies upon the nonspecific adhesion of biomolecules to a surface and a cantilever and, for proteins, the integration of the target protein into a polyprotein. However, this assay results in a low yield of high-quality data, defined as the complete unfolding of the polyprotein. Additionally, nonspecific surface adhesion hinders studies of α-helical proteins, which unfold at low forces and low extensions. Here, we overcame these limitations by merging two developments: (i) a polyprotein with versatile, genetically encoded short peptide tags functionalized via a mechanically robust Hydrazino-Pictet-Spengler ligation and (ii) the efficient site-specific conjugation of biomolecules to PEG-coated surfaces. Heterobifunctional anchoring of this polyprotein construct and DNA via copper-free click chemistry to PEG-coated substrates and a strong but reversible streptavidin-biotin linkage to PEG-coated AFM tips enhanced data quality and throughput. For example, we achieved a 75-fold increase in the yield of high-quality data and repeatedly probed the same individual polyprotein to deduce its dynamic force spectrum in just 2 h. The broader utility of this polyprotein was demonstrated by measuring three diverse target proteins: an α-helical protein (calmodulin), a protein with internal cysteines (rubredoxin), and a computationally designed three-helix bundle (α3D). Indeed, at low loading rates, α3D represents the most mechanically labile protein yet characterized by AFM. Such efficient SMFS studies on a commercial AFM enable the rapid characterization of macromolecular folding over a broader range of proteins and a wider array of experimental conditions (pH, temperature, denaturants). Further, by integrating these enhancements with optical traps, we demonstrate how efficient bioconjugation to otherwise nonstick surfaces can benefit diverse single-molecule studies.

Hidden dynamics in the unfolding of individual bacteriorhodopsin proteins.

Protein folding occurs as a set of transitions between structural states within an energy landscape. An oversimplified view of the folding process emerges when transiently populated states are undetected because of limited instrumental resolution. Using force spectroscopy optimized for 1-microsecond resolution, we reexamined the unfolding of individual bacteriorhodopsin molecules in native lipid bilayers. The experimental data reveal the unfolding pathway in unprecedented detail. Numerous newly detected intermediates-many separated by as few as two or three amino acids-exhibited complex dynamics, including frequent refolding and state occupancies of <10 µs. Equilibrium measurements between such states enabled the folding free-energy landscape to be deduced. These results sharpen the picture of the mechanical unfolding of membrane proteins and, more broadly, enable experimental access to previously obscured protein dynamics.