Live-cell imaging of single integrin tensions with minimal background fluorescence noise.

One powerful method for studying cell mechanobiology is to monitor receptor-mediated forces at the single-molecule level in live cells. Hairpin DNA labeled with a quencher-dye pair has been used as a tension probe (TP) to image cellular forces in real time. The TP emits fluorescence when cellular forces unfold the DNA hairpin and de-quench the dye, thereby converting the force signal into fluorescence. However, when applied to monitor cellular forces at the single-molecule level, the TP often suffers from background fluorescent spots (BFSs) due to nonquenched dyes, which interfere with molecular force imaging and analysis. In this work, we identified that the BFSs are primarily caused by missing quenchers in some TP constructs and surface-adsorbed dye-labeled DNA strands. To address these issues, we developed a double-quencher TP (dqTP) and incorporated Tween-20 treatment during surface preparation. These two simple strategies reduced the BFS level by 10-fold, significantly improving the signal/background ratio for single molecular force imaging. We demonstrated the performance of dqTP by monitoring the temporal dynamics of integrin tensions in platelets and HeLa cells, showing that single integrin tensions remain stable for at least 100 s in wild-type HeLa cells. In contrast, with vinculin knocked out, a subpopulation of integrin tensions, especially at cell peripheral regions, exhibited molecular force fluctuations with an average force duration shorter than 10 s. Overall, this work provides a convenient and practical approach to significantly reduce BFS levels on TP surfaces, offering a nearly false-signal-free platform for monitoring single-molecule forces in live cells.