1. Detection of protein-lipid interaction by flow cytometry

1. Kinetic binding experiments
   1. Dilute phospholipid vesicles in Tyrode's buffer (20 mM HEPES, 150 mM NaCl, 2.7 mM KCl, 1 mM MgCl₂, 0.4 mM NaH₂PO₄, 2.5 mM CaCl₂, 5 mM glucose, 0.5% BSA, pH 7.4) to a concentration of 1 µM and total volume of 250 µL.
   2. Mix fluorescent-labeled coagulation factor X (fX-fd) from step 1 at a concentration of 500 nM with the phospholipid vesicles from step 1.1.1 in a 1:1 ratio (final vesicle concentration 0.5 µM, fX-fd concentration is 250 nM of fX) to a total volume of 500 µL.
   3. Immediately inject the 500 µL of the mixed suspension (~20 min for analysis with a low flowing rate) into the flow cytometer. Use a low flow rate and ensure that the threshold for channel FL2 (excitation 488 nm, emission filter 585/42 nm) is as value 200. Measure the mean fluorescence in channel FL4 (excitation 633 nm, emission filter 660/20) for the fluorescence dye from the Table of Materials.
      NOTE: Choose a cytometer without an autosampler. This will speed up the process of injection of the sample into the measuring cell.
   4. When saturation of binding is achieved (no significant increase in fluorescence within 5 min), rapidly dilute the sample 20-fold with Tyrode's buffer, and monitor the dissociation until baseline fluorescence is reached (complete dissociation) or until a plateau is reached (no significant decrease in fluorescence within 5 min).
      NOTE: As a control, add 10 µM EDTA and monitor complete dissociation for 5 min.