A single vesicle fluorescence microscopy platform to quantify phospholipid scrambling.

Scramblases play important roles in physiology by translocating phospholipids bidirectionally across cell membranes. For example, scrambling facilitated by dimers of the Voltage-Dependent Anion Channel 1 (VDAC1) enables endoplasmic reticulum-derived phospholipids to cross the outer membrane to enter mitochondria. Precise quantification of lipid scrambling, while critical for mechanistic understanding, cannot be obtained from ensemble averaged measurements of reconstituted scramblases. Here, we describe a microscopy platform for high-throughput imaging of single vesicles reconstituted with fluorescently labeled phospholipids and heterogeneously crosslinked VDAC1 dimers. For each vesicle, we quantify size, protein occupancy and scrambling rate. Notably, we find that individual VDAC1 dimers have different activities, ranging from <100 to >10,000 lipids per second. This kinetic heterogeneity, masked in ensemble measurements, reveals that only some dimer interfaces are capable of promoting rapid scrambling, as suggested by molecular dynamics simulations. We extend our analyses to opsin, a monomeric G protein-coupled receptor scramblase, thereby demonstrating the versatility of our platform for quantifying transbilayer lipid transport and exploring its regulation.