Constructing a Magnetic Tweezers to Monitor RNA Translocation at the Single-Molecule Level

Abstract The application of forces and torques on the single molecule level has transformed our understanding of the dynamic properties of biomolecules, but rare intermediates have remained difficult to characterize due to limited throughput. Here, we describe a method that provides a 100-fold improvement in the throughput of force spectroscopy measurements with topological control, which enables routine imaging of 50,000 single molecules and a 100 million reaction cycles in parallel. This improvement enables detection of rare events in the life cycle of the cell. As a demonstration, we characterize the supercoiling dynamics and drug-induced DNA break intermediates of topoisomerases. To rapidly quantify distinct classes of dynamic behaviors and rare events, we developed a software platform with an automated feature classification pipeline. The method and software can be readily adapted for studies of a broad range of complex, multistep enzymatic pathways in which rare intermediates have escaped classification due to limited throughput.

The advent of single-molecule biology has allowed unprecedented insight into the dynamic behavior of biological macromolecules and their complexes. Unexpected properties, masked by the asynchronous behavior of myriads of molecules in bulk experiments, can be revealed; equally importantly, individual members of a molecular population often exhibit distinct features in their properties. Finally, the single-molecule approaches allow us to study the behavior of biological macromolecules under applied tension or torsion; understanding the mechanical properties of these molecules helps us understand how they function in the cell. In this review, we summarize the application of magnetic tweezers (MT) to the study of DNA behavior at the single-molecule level. MT can be conveniently used to stretch DNA and introduce controlled levels of superhelicity into the molecule and to follow to a high definition the action of different types of topoisomerases. Its potential for chromatin studies is also enormous, and we will briefly present our first chromatin results.Key words: single-molecules, chromatin, topoisomerases, magnetic tweezers, force.

AbstractRegulated proteolysis of signaling proteins under mechanical tension enables cells to communicate with their environment in a variety of developmental and physiologic contexts. The role of force in inducing proteolytic sensitivity has been explored using magnetic tweezers at the single-molecule level with bead-tethered assays, but such efforts have been limited by challenges in ensuring that beads are not restrained by multiple tethers. Here, we describe a multiplexed assay for single-molecule proteolysis that overcomes the multiple-tether problem using a flow extension (FLEX) strategy on a microscope equipped with magnetic tweezers. Particle tracking and computational sorting of flow-induced displacements allows assignment of tethered substrates into singly-captured and multiply-tethered bins, with the fraction of fully mobile, single-tethered substrates depending inversely on the concentration of substrate loaded on the coverslip. Computational exclusion of multiply-tethered beads enables robust assessment of on-target proteolysis by the highly specific tobacco etch virus protease and the more promiscuous metalloprotease ADAM17. This method should be generally applicable to a wide range of proteases and readily extensible to robust evaluation of proteolytic sensitivity as a function of applied magnetic force.

This review summarizes the application of magnetic tweezers for the mechanical research of DNA at the single molecule level.

AbstractThe classical “one sequence, one structure, one function” paradigm has shaped much of our intuition of how proteins work inside the cell. Partially due to the insight provided by bulk biochemical assays, individual biomolecules are assumed to behave as identical entities, and their characterization relies on ensemble averages that flatten any conformational diversity into a unique phenotype. While the emergence of single-molecule techniques opened the gates to interrogating individual molecules, technical shortcomings typically limit the duration of these measurements to a few minutes, which prevents to completely characterize a protein individual and, hence, to capture the heterogeneity among molecular populations. Here, we introduce a magnetic tweezers design, which showcases enhanced stability and resolution that allows us to measure the folding dynamics of a single protein during several uninterrupted days with a high temporal and spatial resolution. Thanks to this instrumental development, we do a complete characterization of two proteins with a very different force-response: the talin R3IVVI domain and protein L. Days-long recordings on the same single molecule accumulate several thousands of folding transitions sampled with sub-ms resolution, which allows us to reconstruct their free energy landscapes and describe how they evolve with force. By mapping the nanomechanical identity of many different protein individuals, we directly capture their molecular diversity as a quantifiable dispersion on their force response and folding kinetics. Our instrumental development offers a new tool for profiling individual molecules, opening the gates to the characterization of biomolecular heterogeneity.

Histone H1 binds to linker DNA between nucleosomes, but the dynamics and biological ramifications of this interaction remain poorly understood. We performed single-molecule experiments using magnetic tweezers to determine the effects of H1 on naked DNA in buffer or during chromatin assembly in Xenopus egg extracts. In buffer, nanomolar concentrations of H1 induce bending and looping of naked DNA at stretching forces below 0.6 pN, effects that can be reversed with 2.7-pN force or in 200 mM monovalent salt concentrations. Consecutive tens-of-nanometer bending events suggest that H1 binds to naked DNA in buffer at high stoichiometries. In egg extracts, single DNA molecules assemble into nucleosomes and undergo rapid compaction. Histone H1 at endogenous physiological concentrations increases the DNA compaction rate during chromatin assembly under 2-pN force and decreases it during disassembly under 5-pN force. In egg cytoplasm, histone H1 protects sperm nuclei undergoing genome-wide decondensation and chromatin assembly from becoming abnormally stretched or fragmented due to astral microtubule pulling forces. These results reveal functional ramifications of H1 binding to DNA at the single-molecule level and suggest an important physiological role for H1 in compacting DNA under force and during chromatin assembly.

Mechanochemical analysis of DNA and RNA double helices at the single-molecule level by force- and torque-measuring optical and magnetic tweezers.

DNA supercoiling is a key regulatory mechanism that orchestrates DNA readout, recombination, and genome maintenance. DNA-binding proteins often mediate these processes by bringing two distant DNA sites together, thereby inducing (transient) topological domains. In order to understand the dynamics and molecular architecture of protein induced topological domains in DNA, quantitative and time-resolved approaches are required. Here we present a methodology to determine the size and dynamics of topological domains in supercoiled DNA in real-time and at the single molecule level. Our approach is based on quantifying the extension fluctuations -in addition to the mean extension- of supercoiled DNA in magnetic tweezers. Using a combination of high-speed magnetic tweezers experiments, Monte Carlo simulations, and analytical theory, we map out the dependence of DNA extension fluctuations as a function of supercoiling density and external force. We find that in the plectonemic regime the extension variance increases linearly with increasing supercoiling density and show how this enables us to determine the formation and size of topological domains. In addition, we demonstrate how transient (partial) dissociation of DNA bridging proteins results in dynamic sampling of different topological states, which allows us to deduce the torsional stiffness of the plectonemic state and the kinetics of protein-plectoneme interactions. We expect our approach to enable quantification of the dynamics and reaction pathways of DNA processing enzymes and motor proteins, in the context of physiologically relevant forces and supercoiling densities.