Samples:
1) Cultured cells: Cells may be grown on a 12mm round coverslips and stained in the wells of a 24-well plate. Alternatively cells can be grown in a petri dish, in which a hole has been made and a coverslip is glued in (Mat-Tek Corp. 200 Homer Ave, Ashland MA 01721, (800) 834-9018). Finally, now that we have an upright confocal microscope cells maybe grown on plastic and viewed live with the Zeiss LSM 510 Meta using dipping objecives.
2) Sections of fixed tissues: Sections up to 200 µm thick made with a Vibrotome can be viewed on a 2-photon microscope. For standard confocal (single photon), 50-100 µm should be max.
3) Thin sections: These should be made as usual for histological staining. After attaching to glass slide (coated with poly-L-lysine or purchased charged slides: Vector Labs), they must be fixed.

Fixation: Since different fixatives can have various effects on cytoskeletal structures or on proteins by cross-linking, several methods of fixation should be tested for each antibody developed:
1) 3.7% paraformaldehyde (PFA) in PBS for 15 minutes at room temperature followed by 0.2% Triton X100 in PBS (5 min) or 100 µM digitonin (5-10 min) permeabilization. This is the fix of choice if it works with your antibodies, because the cytoskeleton is best preserved. (Permeabilization may not be necessary in very thin sections.)
2) Absolute methanol at -20°C for 6 minutes; flood with PBS-0.5% BSA to prevent drying when incubation is finished.
3) Absolute ethanol at room temperature for 30 seconds; flood with PBS-0.5% BSA to prevent drying when incubation is finished.

Fluorescence staining:
1) Wash the cells several times in PBS-0.5% BSA. (I just use a squirt bottle and thoroughly flush the slide several times or fill and dump in the case of cells grown on coverslips and incubated in 24-well plates.)
2) Incubate with primary antibodies in PBS-0.5% BSA for 1 hour at room temperature or a half hour at 37°C. I usually only make enough working dilution for each experiment and don’t reuse it or save left over. The total amount I use for 24-well plates or for circles drawn on slides is 10µl. But I don’t measure less than a microliter of stock; so if it should be diluted 1:200, I make 200 µl. If there are two antibodies I mix them first, then place the solution on the cells. Whether on a slide or on coverslips in wells this incubation should be done in a humid chamber. This can be accomplished by placing the slide on toothpicks on a damp paper towel in a plastic box of some sort, or placing the damp paper towel in the lid of the 24-well plate.
3) After incubation, wash again in PBS, and add fluorescent secondary antibodies (at this point I add the direct labeling reagents, eg DAPI or phalloidin, as well), followed again by washing. Only secondary antibody should be incubated on one sample to control for nonspecific reaction. Incubate and wash again as above.

Mounting: They are embedded in an anti-fade media (available from Molecular Probes or Vector labs). There are of course many variations on this media. It is only important that the issue of fading be addressed somehow. I will often rinse my cells for preparation once in deionized water before mounting too. This simply rinses off any salt residue, but is not necessary for viewing. Thick sections of 60-200 µm may be stored in a PBS a solution containing 2% DABCO instead of mounting.