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.
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...
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.
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