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.
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.
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.
Quantifying the energy landscape underlying protein-ligand interactions leads to an enhanced understandingo fm olecular recognition.Apowerful yet accessible single-molecule technique is atomicf orce microscopy (AFM)-based force spectroscopy,w hich generally yields the zero-force dissociation rate constant( k off )a nd the distance to the transitions tate (Dx°). 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 yieldedt he height of the transition state (DG°= 6.3 AE 0.2 kcal mol À1)a nd the shape of the energy barrier at the transition state (linear-cubic) in addition to the traditional parameters [k off (= 4 AE 0.1 10 À4 s À1)a nd Dx°(= 8.3 AE 0.1 )]. We expect this automated and relativelyr apid assay to provide am ore complete energy landscape description of proteinligand interactions and, more broadly,t he diverse systems studied by AFM-based force spectroscopy.Molecular recognition between proteins and ligands is fundamentalt ob iology.C orrect recognition of antigens by antibodies, substrates by enzymes, and ligands by receptors is essential to most biological processes. In addition, the ability to custom-design proteins with precise and selectivem olecular recognition for at arget molecule would enablet he development of biosensors for aw idea rray of biological and medical applications.Characterizing the strength of natural and computationally designed protein-ligand interactions is usually done in bulk assays,y ielding measurements of the dissociation constant (K D ). For instance, DIG10.3, which binds the steroid digoxigenin (Dig), is the first computationally designed protein to achievea picomolarl evel K D to its target ligand.[1] Indeed, DIG10.3e xhibits an affinity that rivals that of anti-Dig antibodies.[2] Molecular details of the bound state are provided by structurals tudies (e.g. X-ray crystallography) and have confirmed the computationally predicted binding mode.[1] However,e xperimentald etermination of the process of dissociation remains elusive. Hence, understandingo fp rotein-ligand interactions would benefitf rom an expanded description of the free-energy landscape that governs dissociation, including the height( DG°) and distance (Dx°)t ot he transition state along with the shape of the free-energy barrier at the transition state.Single-molecule force spectroscopy (SMFS)i sapowerful technique to characterize protein-ligand interactions. [3][4][5][6][7] In such assays, af orce applieda cross the protein-ligand interaction promotes detachment. The resultingd ata, often taken with an atomic force microscope (AFM)o ver ar ange of stretching velocities and thereby loading rates (@F/@t), [7] yields insight into the energy landscape underlying the proteinligand interaction projected onto the stretching axis. [8] Standard analysis uses the Bell-Evans model,w hich predicts al inear relationship between the most probable rupture force and log(@F/@t)and...
The Front Cover visualizes the concept of using single‐molecule force spectroscopy based on atomic force microscopy to better quantify the energy landscape underlying protein–ligand interactions. We introduce an enhanced AFM assay and apply it to probe the binding of a computationally designed protein to its target ligand. More information can be found in the Communication by W. J. Van Patten et al. on page 19 in Issue 1, 2018 (DOI: 10.1002/cphc.201701147).
The front cover artwork is provided by the groups of David Baker (Univ. of Washington and HHMI) and Thomas Perkins (JILA, NIST and Univ. of Colorado). The image shows an AFM cantilever pulling on a computationally designed protein bound to digoxigenin attached to the end of a short DNA molecule with a conceptual free‐energy landscape as background. Read the full text of the article at 10.1002/cphc.201701147.
Conclusion: Acute knockdown of desmin in isolated adult rat ventricular myocytes reveals previously unidentified structural and functional roles of this IF. Further study must be done to identify how desmin upregulation affects disease and other roles desmin may play in mechanotransduction.
Programmed À1 frameshifting (À1 PRF) is an important biological process for the modification of gene expression enabled by the presence of RNA secondary and tertiary structures. However, the mechanism for this process is still under active study. Here, we investigate the role of mechanical force in À1 PRF by characterizing mechanically induced folding and unfolding of RNA pseudoknots using an enhanced atomic force microscopy (AFM) based singlemolecule force spectroscopy (SMFS) assay featuring $10 ms resolution. The pioneering SMFS study of RNA psuedoknots associated with À1 PRF used a custom-built optical trap. Unexpectedly, this study indicated PRF efficiency was correlated with the presence of alternative folding pathways (rather than with average unfolding force), indicating a complex role of RNA psuedoknots in À1 PRF involving folding dynamics. Here, we found a new folding intermediate in an RNA pseudoknot associated with the sugarcane yellow leaf virus (ScYLV) that was not observed in the original optical-trapping based assay. We speculate that the shorter linkers and stiffer force probe in our AFM assay (relative to an optical trap) are the primary reasons for this enhanced state resolution. Our initial measurements of contour length and folding kinetics indicate this folding intermediate is an RNA hairpin that is part of the overall pseudoknot structure. We observed this intermediate every time the pseudoknot folds, indicating that this intermediate is obligate for folding. Overall, our results indicate that the folding dynamics of RNA pseudoknots are significantly more complex than previously observed. Because of the role of the folding dynamics of RNA pseudoknots in À1 PRF, we expect these new insights into RNA pseudoknot folding dynamics will provide a deeper understanding into the mechanisms of À1 PRF.
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