Take out the slides to a slide holder and warm slides on a 37°C heater for 5-10 min. Grouping: usually maximum two antibodies (with secondary antibody red + green) + nuclear staining for one slide. Set negative control for each secondary antibody to detect any non-specific background.

Label slides with pencil and circle section with Pap Pen (hydrophobic marker). Move the slides into a humidified chamber. Slides should remain in humidified chamber for all the following incubation steps.

Fixation: Add one or two drops of 4% parformaldehyde in PBS on the tissue for 15-30 min.

Wash with 0.5% Tween 20 in PBS (PBST) , 10 min x 2.

Block with Blocking buffer: 10% host serum + 2% FBS in 0.5% PBST. Incubate 1 hour at room temperature or 4°C overnight.

Incubate with primary antibody: 1-10 μg/mL of purified antibody or 1:100 to 1:1000 dilution of anti-serum in 2% host serum + 2% FBS 0.5% PBST buffer overnight at 4°C. Normally start with a dilution at 1:100. For color-conjugated FC antibodies used in immunofluorescence, start at 1:50.

Wash with 0.5% PBST, 10 min x 2.

Incubate with secondary antibody: 1-10 μg/mL of purified antibody or 1:100-1:1000 of anti-serum in Blocking buffer (normally start at 1:200). Incubate 1 h at room temperature ( AVOID LIGHT ).

Wash with PBST, 10 min x 3.

Incubate the slides in 0.1 μg/mL DAPI for 5 min.

Wash with PBST twice before mounting.

Clean the slides.

Mount the slides with coverslips using anti-fade mounting medium.

Seal the edges of the coverslip with nail polish and let it dry.