Transmission Electron Microscopic Imaging of Synaptic Vesicle Endocytosis in Mouse Hippocampal Neurons

Begin by treating mouse hippocampal neurons with a high-potassium buffer containing horseradish peroxidase or HRP.

High potassium depolarizes presynaptic neurons, triggering synaptic vesicle exocytosis. The neurons then retrieve the synaptic vesicle membranes through endocytosis, by incorporating HRP into the newly formed vesicles. Fix the neurons with glutaraldehyde and wash to remove excess glutaraldehyde.

Add hydrogen peroxide and a chromogenic substrate, which is oxidized by HRP to form electron-dense precipitates. Wash to remove any unreacted substrate.

Treat with osmium tetroxide to enhance membrane contrast, then wash off the excess reagent. Next, add uranyl acetate to enhance organelle contrast.

Dehydrate the neurons using increasing ethanol concentrations and embed them in resin.

Microscopically identify cell-dense regions and excise them. Then, prepare ultra-thin sections.

Transfer a section onto a copper grid and counterstain them with contrast agents.

Using transmission electron microscopy, vesicles containing HRP appear as electron-dense structures.

Stimulate the hippocampal neuron culture by adding 1.5 milliliters of high-potassium stimulation solution to each well at room temperature for 90 seconds. In this step, prepare a 0.1 molar sodium cacodylate buffer at pH 7.4. Fix the cells with 4% glutaraldehyde in 0.1 molar sodium cacodylate buffer for at least one hour at room temperature.

Then, wash the cells three times with 0.1 molar sodium cacodylate buffer each time for seven minutes. Following that, prepare DAB solution and filter it with the 0.22 micrometer filter. Next, apply 1.5 milliliters of DAB solution to the cells for 30 minutes at 37 degrees Celsius.

Then, wash the cells three times with 0.1 molar sodium cacodylate buffer each time for seven minutes. As post-fixation, incubate the neurons with 1.5 milliliters of 1% osmium tetroxide in 0.1 molar sodium cacodylate buffer for one hour at four degrees Celsius.

Afterward, wash the neurons three times with 1.5 milliliters of 0.1 molar sodium cacodylate buffer each time for seven minutes. Subsequently, prepare 0.1 molar sodium acetate buffer with sodium acetate and glacial acetic acid. Wash the cells three times with 1.5 milliliters of 0.1 molar acetate buffer each time for seven minutes.

After that, incubate the cells with 1.5 milliliters of 1% uranyl acetate in 0.1 molar acetate buffer for 1 hour at four degrees Celsius. Then, wash the cells three times with 1.5 milliliters of 0.1 molar acetate buffer for seven minutes each time. In a fume hood, dehydrate the neuronal culture with single 1.5 millileter washes of 50%, 70%, and 90% ethanol for seven minutes each wash, followed by three washes with 1.5 milliliters of 100% ethanol for seven minutes each time.

Next, create the epoxy resin and mix thoroughly and store it under vacuum to remove air bubbles. Infiltrate the sample by replacing the ethanol with 50% epoxy resin in ethanol on a shaker for 30 minutes at room temperature, then 70% epoxy resin and ethanol for 30 minutes at room temperature on a shaker.

Switch the epoxy resin solution with 100% epoxy resin and incubate for 10 minutes at 50 degrees Celsius. Perform two exchanges of fresh 100% epoxy resin. Each exchange is incubated for one hour at 50 degrees Celsius. Then, add fresh 100% epoxy resin and allow it to harden at 50 degrees Celsius overnight and then at 60 degrees Celsius for over 36 hours.

Afterward, remove each sample from the multi-well plate with the jeweler's handsaw. Using an inverted light microscope, select the regions of interest that have dense concentrations of cells. Then, cut the blocks of 4 by 5 by 8 millimeters cubed for sectioning. Afterward, counterstain the sections by submersion with 1% aqueous uranyl acetate for 15 minutes and then 3% aqueous-lead citrate for 5 minutes to improve the contrast of the samples.