You could preload the cells with a fluorescent dextran (e.g., 70 kDaltons) by pinocytosis overnight and then rinse well. You could then measure release of the dextran into the medium (or loss from the cells) by fluorescence measurements.If you are imaging live cells and use a pH-depepndent fluorophore(e.g., FITC), you can see a puff of fluorescence as the FITC-dextran in the acidic lysoosmes is released into the neutral pH medium. The dextran will rapidly diffuse away. This is probably most easily seen with TIRF microscopy.

As an alternative, you could measure lysosomal enzyme release using a fluorogenic substrate, but you will need to correct for constitutive secretion in the absence of your activation of Ca-dependent release.

Hi Magnus, 

There is way to quantify the exocytosis in cultured cells live. FM4-64 dyes are used for this purpose. When incubated with cells, these dyes fluoresces only on internalization into lysosomes. It takes approximately 20-30 min for internalization (endocytosis). After internalization you can wash the cells and add fresh medium containing calcium. Exocytosis occurs and it this process can be imaged under fluorescence microscope. Quantification of captured images after frequent washes for upto 30 min will give you the information on magnitude of exocytosis. 

good luck

Rajesh

Thank you for your suggestions! Is it known whether or not the lysosomal membrane fuse with the plasma membrane during lysosomal exocytosis? If so, could levels of lysosomal membrane proteins, such as Lamp-2, in the plasma membrane serve as a marker for the process.

The lysosome membranes do fuse with the plasma membrane. However, the LAMP proteins are rapidly endocytosed back into the cell.