Single-molecule manipulation studies have revealed that doublestranded DNA undergoes a structural transition when subjected to tension. At forces that depend on the attachment geometry of the DNA (65 pN or 110 pN), it elongates Ϸ1.7-fold and its elastic properties change dramatically. The nature of this overstretched DNA has been under debate. In one model, the DNA cooperatively unwinds, while base pairing remains intact. In a competing model, the hydrogen bonds between base pairs break and two single DNA strands are formed, comparable to thermal DNA melting. Here, we resolve the structural basis of DNA overstretching using a combination of fluorescence microscopy, optical tweezers, and microfluidics. In DNA molecules undergoing the transition, we visualize double-and single-stranded segments using specific fluorescent labels. Our data directly demonstrate that overstretching comprises a gradual conversion from double-stranded to singlestranded DNA, irrespective of the attachment geometry. We found that these conversions favorably initiate from nicks or free DNA ends. These discontinuities in the phosphodiester backbone serve as energetically favorable nucleation points for melting. When both DNA strands are intact and no nicks or free ends are present, the overstretching force increases from 65 to 110 pN and melting initiates throughout the molecule, comparable to thermal melting. These results provide unique insights in the thermodynamics of DNA and DNA-protein interactions.DNA melting ͉ fluorescence microscopy ͉ optical trapping ͉ single-molecule techniques ͉ single-stranded DNA

The central catalyst in eukaryotic ATP-dependent homologous recombination consists of RAD51 proteins, polymerized around single-stranded DNA. This nucleoprotein filament recognizes a homologous duplex DNA segment and invades it1,2. After strand exchange, the nucleoprotein filament should disassemble in order for the recombination process to complete3. The molecular mechanism of RAD51 filament disassembly is poorly understood. Here, we have combined optical tweezers with single-molecule fluorescence microscopy and microfluidics4,5 to reveal that disassembly results from the interplay between ATP hydrolysis and release of the tension stored in the nucleoprotein filament. Applying external tension to the DNA, we found that disassembly slows down and can even be stalled. We quantified the fluorescence of RAD51 patches and found that disassembly occurs in bursts interspersed by long pauses. Upon relaxation of a stalled complex, pauses were suppressed resulting in a large burst. These results imply that tension-dependent disassembly takes place only from filament ends, after tension-independent ATP hydrolysis. This integrative single-molecule approach allowed us to dissect the mechanism of this key homologous recombination reaction step, which in turn clarifies how disassembly can be influenced by accessory proteins.

The DNA strand-exchange reactions defining homologous recombination involve transient, nonuniform allosteric interactions between recombinase proteins and their DNA substrates. To study these mechanistic aspects of homologous recombination, we produced functional fluorescent human RAD51 recombinase and visualized recombinase interactions with single DNA molecules in both static and dynamic conditions. We observe that RAD51 nucleates filament formation at multiple sites on double-stranded DNA. This avid nucleation results in multiple RAD51 filament segments along a DNA molecule. Analysis of fluorescent filament patch size and filament kinks from scanning force microscopy (SFM) images indicate nucleation occurs minimally once every 500 bp. Filament segments did not rearrange along DNA, indicating tight association of the ATP-bound protein. The kinetics of filament disassembly was defined by activating ATP hydrolysis and following individual filaments in real time.

Many biological processes involve enzymes moving along DNA. Such motion might be impeded by DNA-bound proteins or DNA supercoils. Current techniques are incapable of directly measuring forces that such 'roadblocks' might impose. We constructed a setup with four independently moveable optical traps, allowing us to manipulate two DNA molecules held between beads. By tightly wrapping one DNA around the other, we created a probe that can be scanned along the contour of the second DNA. We found that friction between the two polymers remains below 1 pN. Upon encountering DNA-bound proteins substantial friction forces are measured, allowing accurate localization of protein positions. Furthermore, these proteins remained associated at low probe tensions but could be driven off using forces greater than 20 pN. Finally, the full control of the orientation of two DNA molecules opens a wide range of experiments on proteins interacting with multiple DNA regions.