Abstract
Functional activities of many transmembrane proteins are controlled by their endocytosis. One of the most studied experimental models is the epidermal growth factor (EGF) receptor (EGFR). However, endocytic trafficking of EGFR has been predominantly analyzed using labeled EGF, whereas quantitative analyses of the endocytosis of the receptor itself have been sparse. The fluorescence microscopy methods described here are designed to directly quantify EGFR internalization in living cells without labeled EGFR ligands or antibodies. These methods utilize an engineered EGFR chimera in which the fluorogen activating protein (FAP) is fused to the receptor extracellular domain (FAP-EGFR). Binding of malachite green (MG) based dyes to FAP results in a strong far-red fluorescence of MG, thus efficiently labeling FAP-EGFR. In particular, binding of the cell impermeant MG-Bis-SA dye to FAP produces the pH-sensitive dual-excitation fluorescence, which allows differentiation of the cell-surface and internalized pools of FAP-EGFR. Two modifications of the methodology are described: 1) single-cell three-dimensional confocal imaging; and 2) high-throughput assay in multi-well plates. These methodologies can be adopted to study endocytosis of any other transmembrane protein extracellularly tagged with FAP.