The measurement of dissociation constant (𝐾𝑑) between a protein and a ligand reveals their binding affinity. 𝐾𝑑 is an equilibrium constant widely used in various fields. In pharmacology, it describes the affinity of drugs to their targets; in enzymology, it characterizes the interactions between enzymes and substrates; and in cell signaling and immunology, it describes the affinity between ligands and receptors. In the context of proteins-nucleic acids interactions, 𝐾𝑑 is widely used to quantify the affinity between DNA and proteins such as transcription factors, DNA binding enzymes, or structural proteins like nucleosomes. 𝐾𝑑 is used to distinguish between sequence-specific and non-specific binding, thereby describing the functional role and the mechanism of action of proteins that participate in regulation of gene expression. However, 𝐾𝑑 characterizes an equilibrium state of the system, whereas genome processing is highly dynamic, driven by the interplay of numerous structural, regulatory, and motor proteins. Genome processing occurs outside of equilibrium, rendering 𝐾𝑑 a less reliable predictor of the outcome of dynamic interactions, such as competition for binding site, competition between genome compaction and processing, and encounters between motor proteins and DNA-bound proteins. A more suitable description can be achieved by using constants that describe the kinetics of complex formation and dissociation. For instance, the association rate constant (𝑘𝑜𝑛), describes the speed of protein association with their binding sites, whereas the dissociation rate constant (𝑘𝑜𝑓𝑓) describes the binding stability of a protein on its DNA binding site. A protein with low 𝑘𝑜𝑓𝑓 would not be easily displaced from DNA and could function as a transcription or replication termination factor. Whereas, proteins involved in dynamic processes would be expected to have high 𝑘𝑜𝑛 to facilitate rapid binding and high 𝑘𝑜𝑓𝑓 to promote their displacement. Equally important is the characterization of mutants. For instance, a mutation in a transcription termination factor, as seen in mitochondrial transcription termination factor (MTERF1), results in the synthesis of longer mRNA. Therefore, understanding whether the mutation affects the probability of binding (𝑘𝑜𝑛) or the strength of binding (𝑘𝑜𝑓𝑓) is of primary importance. In one case, MTERF1 will be present on the termination site less often, yet the protein will remain functional. In the second case, iv MTERF1 would be present on the termination site as often as the wild-type, but it would be easily displaced by the mitochondrial RNA-polymerase (RNAP). However, the methods used to characterize kinetic rate constants in proteinDNA interactions are not optimized for specific requirements of the field. One of the primary methods for measuring kinetic rate constants is surface plasmon resonance, widely recognized under its commercial name Biacore. Although this technique is well established, it fails to capture the very fast binding kinetics and low unbinding rates typical for strong interactions between proteins and DNA. It also lacks the ability to differentiate between specific and non-specific interactions and does not address sample heterogeneity. In addition, this approach requires arrays of densely immobilized short oligonucleotides as a substrate, which can induce mass transport and surfaceassociated limitations. Other bulk methods like fluorescence anisotropy, do not have surface-associated limitations but require protein and DNA fluorescent labeling. Single-molecule fluorescence, although providing a direct read-out of binding and unbinding events, poses experimental challenges due to labeling requirements. Additionally, it requires extra controls to distinguish between binding events and artifacts, as well as unbinding events and the blinking or bleaching of fluorophores. All this together, underscores the need for new experimental approaches capable of addressing the potential heterogeneity of binding and unbinding kinetics in a sample, in a label-free manner, with sensitivity comparable to current state-of-the-art techniques. To address this need, we introduce a hairpin assay in combination with magnetic tweezers. This approach not only fulfills the current requirements but also offers advantages, including zero dead-time needed to probe very fast binding kinetics, force-assisted unbinding that can speed up unbinding in case of proteins with very low 𝑘𝑜𝑓𝑓, and low experimental volumes common for single-molecule experimentation. Additionally, our approach allows for the probing of protein unbinding in the context of dynamic unwinding of double-stranded DNA. Increasing evidence suggests that termination factors sense the directionality of approaching motor proteins, making our method invaluable for understanding binding strength in the context of dynamic protein behavior. Here, we describe a method to probe the binding of macromolecules to single- and double-stranded DNA, along with an extensive characterization of the method using sgRNA and Cas9 to describe binding to both single- and double-stranded DNA. Using MTERF1, we illustrate different protein binding times depending on the direction of DNA unwinding, and we describe the workflow required to address protein unbinding from DNA. We also demonstrate how sensitive the measurements of unbinding rates are to force variability, and describe new force calibration routines and approaches to improve the stability of magnetic tweezers. We believe that with the increased usage of magnetic tweezers and other highthroughput force spectroscopy instruments, such as acoustic force spectroscopy and centrifugal force microscope, our work will make a valuable contribution to the field of single-molecule characterization of protein-DNA interactions.
Eugeniu Ostrofet (Thu,) studied this question.