The following protocol has been adapted from one from Larry Zipursky's lab. It detects antigen in retinas that have been sectioned in plastic.
Cut heads in half, taking care not to place excessive pressure on the head. Dip briefly in 70% ethanol and immerse in 4% formaldehyde in PEM buffer. Fix at room temperature for 60 minutes.
Transfer to BSS and wash once. Dissect retinas at this point. Cut along the edge of the lens and remove retina and cornea from the rest of the head. Remove with forceps or hook, a thin pigmented layer of cells located between the retina and the lamina. Then remove the cornea from the retina by inserting a hook between the retina and cornea. Take care as the tissue is soft.
Wash three times for 10 minutes each in BSS at room temperature. Permeabilize 30 minutes in BSSDT at room temperature.
Incubate overnight at 4oC (shaking) in primary antibody diluted in BSSDT. Wash four times for 10 minutes each in BSSDT.
Incubate two hours in secondary antibody diluted in BSSDT. Wash four times for 10 minutes each in PBT. The last wash is at room temperature.
Incubate in 0.5 mg/ml DAB, 0.01% NiCl2 in PBT for 5 minutes at room temperature. Transfer to 0.5 mg/ml DAB, 0.01% NiCl2, 0.003% H2O2 in PBT and develop for 5 to 15 minutes at room temperature.
Wash twice in PBT for 5 minutes each and once in PBS for 5 minutes.
Dehydrate and embed in Durcapan resin as normally done for retinal sectioning. Embed retinas in BEEM or gelatin capsules with the outer face of the retina facing downwards.
Cut 2 mm sections. No post-staining is required. Examine sections using Nomarski or phase contrast optics.
2.21 g NaCl
3.98 g KCl
3.07 g MgSO4.7H2O
0.74 g CaCl2.2H2O
1.79 g Tricine
3.60 g glucose
17.12 g sucrose
2.00 g BSA
add H2O to 1 liter, adjust pH to 6.95, filter sterilize. I make a 2X stock solution.
Add Sodium Deoxycholate to 0.3% and Triton X100 to 0.3% final concentrations in BSS.
Add Triton X100 to 0.1% final concentration in PBS.