This article describes a protocol for analyzing synaptic vesicle recycling in cultured neurons using fluorescent-tagged proteins. The method involves depolarization, antibody binding, and optical analysis to visualize recycled vesicles.
Begin with a coverslip containing cultured neurons expressing synaptic vesicle membrane proteins, including fluorescent-tagged synaptophysin and synaptotagmin-1.
Remove the medium, add a mix of depolarization buffer and primary antibodies, then incubate.
The buffer ions enter the cells, stimulating the fusion of synaptic vesicles with the membrane, which exposes synaptotagmin-1.
The primary antibodies selectively bind to the exposed synaptotagmin-1.
Discard the buffer and wash with the medium to remove unbound antibodies.
Introduce fresh medium and incubate to facilitate membrane internalization, forming recycled vesicles with labeled synaptophysin and antibody-bound synaptotagmin-1.
Remove the medium, fix the cells, and wash.
Introduce a detergent-based blocking buffer with fluorophore-coupled secondary antibodies.
The detergent permeabilizes the cells, allowing the secondary antibodies to interact with antibody-bound synaptotagmin-1.
Lift the coverslip, wash it with water, and dry it.
Place the coverslip face-down on a slide with an embedding medium for optical analysis of the dual-labeled recycled vesicles.
Prewarm 600 microliters of 1x depolarization buffer and 10 milliliters of cell culture medium to 37 degrees Celsius in the water bath. Add 1 microliter of mouse anti-SYT1 antibody to the 1x depolarization buffer and vortex for 10 seconds. After removing and discarding the cell culture medium from the cells, add 200 microliters of the depolarization antibody mix to each well.
Incubate the plate for 5 minutes at 37 degrees Celsius and 5% CO2 in the incubator. Remove and discard the depolarization antibody mix and wash the cells three times with cell culture medium. Add 1 milliliter of cell culture medium to each well and incubate for 30 seconds at room temperature.
Then, remove 750 microliters of medium and add the same amount of fresh medium. Repeat this process three times. Next, remove and discard the cell culture medium and add 300 microliters of 4% PFA in 1x PBS.
Finally, incubate for 20 minutes at 4 degrees Celsius before washing three times for 5 minutes each with 1x PBS. Dilute the secondary fluorophore-coupled antibody in 200 microliters of blocking buffer for each well at a dilution of 1 to 1,000. Remove and discard the 1x PBS from each well containing coverslips with the cultured neurons.
Add 200 microliters of blocking buffer antibody mix to each well, and incubate for 60 minutes at room temperature. After incubation, wash the cells three times for 5 minutes with 1 milliliter of 1x PBS.
Next, add a 7-microliter drop of embedding medium onto the microscope slide. Remove the coverslip from the 24-well plate by lifting it with a cannula and grabbing it with forceps. Dip the coverslip into distilled water to remove the PBS, and dry it carefully by touching one edge to a soft tissue.
Flip the coverslip onto the embedding medium droplet so that the surface carrying the cells faces the microscope slide, thereby embedding the cells into the embedding medium. Leave the slides to dry under the hood for one to two hours, covering them to avoid light exposure. Store the dried slides in a microscope slide box at 4 degrees Celsius.
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