Epi-fluorescence microscopy.

5.

Sample Immuno-Fluorescence Staining Protocol

Fix cells grown in vitro on coverslips by incubating as follows: Presynaptic Proteins: 4% paraformaldehyde (PFA)/4% sucrose in phosphate buffered saline (PBS) for 15 min at room temperature. Postsynaptic Density Proteins: methanol at –20°C for 20 min. PSD and EGFP Fluorescence: 1% PFA/4% sucrose in PBS for 3 min at room temperature, followed by methanol at –20°C for 10 min. Rinse three times with PBS. Permeabilize cells with 0.2% Triton X-100 for 5 min at room temperature. Rinse three times with PBS. Place coverslips in a humid environment to prevent them from drying out. To create a humid environment, wet Kimwipes can be placed in the incubation chamber. Incubate with 100 μL of 20% goat serum at room temperature for 1 h to block nonspecific binding. Dilute primary antibody in 5% goat serum at a dilution of 1:100. Note: This is a good starting concentration but lower or higher concentrations may be fine depending on the antibody. Always test a number of concentrations and use the lowest concentration that gives good specific binding and little nonspecific binding. Incubate the coverslips with 100 μL of primary antibody for 1 h at room temperature or overnight at 4°C. Rinse three times with PBS. Dilute a fluorophore-conjugated secondary antibody 1:500 in 5% goat serum, and store in the dark. Again, the concentration of secondary antibody should be tested and the lowest concentration that gives good specific and minimal nonspecific labeling should be used. Incubate the coverslip with 100 μL of secondary antibody for 1 h at room temperature. Do not incubate with secondary antibodies overnight, as it typically increases nonspecific binding without any significant improvement in specific binding. This may not be the case for thick tissue samples where it may take time for the secondary antibody to penetrate the tissue. Rinse three times with PBS. Place ~10 μL of mounting media containing an anti-fade agent (e.g., Aquamount; ThermoFisher) on a microscope slide and invert the coverslip onto the slide. Use a cotton swab to press down gently on the coverslip and direct any air bubbles to the edges. Coverslips can be sealed with clear nail polish to prevent evaporation. However, there is evidence that the solvents in nail polish can alter the specimen integrity or the fluorescence of certain dyes. A more benign sealant is VALAP, which is a 1:1:1 mixture of Vaseline, lanolin, and paraf fin. This mixture can be made by melting the three components in a beaker on a hot plate, letting it cool and then cutting it into small blocks. These blocks can be melted and dripped onto the edges of coverslips using a glue gun or soldering iron. A third option is to use a product called Twinseal available from Picodent, Wipperfurth, Germany. The yellow and blue components are mixed 1:1 and spread around the coverslip and left for 5 minutes to harden. In general, a mounting media that cures and hardens is the best for specimen preservation (e.g., Shandon Immu-Mount-ThermoFisher; CytoSeal-Edmund Scientific; Prolong Gold-Invitrogen). Slides can be kept for weeks or months when stored in the dark at 4°C. Neurons were stained for synapses with an SV2 ( Fig. 5a ) primary antibody and AlexaFluor® 488 (Life Technologies) secondary antibody. Filamentous actin was stained with phalloidin-TRITC ( Fig. 5b ). Phalloidin-TRITC was incubated in the same step as the primary anti-SV2 antibody after goat serum blocking. Phalloidin is commonly used to visualize dendritic spines since filamentous actin is abundant in these structures ( Fig. 5b ) ( 51 , 52 ). Note: phalloidin staining does not work well with methanol fixation. For live cell work, transfection with plasmids for the membrane localized GAP-GFP protein conjugate (the palmitoylation and membrane targeting domain of neuromodulin (GAP-43)) provides a nice outline of dendrites and axons ( 53 ).