Immunocytochemistry (ICC)

10/24/00

1. You will need to thaw 8% paraformaldehyde and a tube of goat serum.

2. In the hood, remove excess medium in the wells of the plate until (for a 24 well plate) there is 0.5mL per well.

3. FIX: Add 0.5mL/well (an equal volume to the well volume) of 8% paraformaldehyde to each well. This gives a final concentration of 4%. Fix for 15 minutes at room temp.

4. While the cells are fixing, make blocking solution:

8.4mL 1X PBS pH 7.4

1.6mL goat serum (final = 16%)

100uL 20% tritonX-100 (final = 0.2%)

5. Suck off the paraformaldehyde with your pipetter and place in a tube. Dispose of paraform. in the bottle under the hood near the tc room. Do not put paraformaldehyde down the sink.

6. Wash the cells with 1X PBS, 0.5mL/well. You may now aspirate the solutions off the cells.

7. BLOCK: Add 0.35mL blocking solution/well to the cells. Leave at room temp for 60 minutes.

8. Meanwhile, make the dilutions of primary antibody in blocking solution. Each antibody has its own dilution, so check how much you should use each time you stain cells. You will need 50uL for each coverslip, so calculate how much of each dilution you will need total.

anti-myc epitope (9E10 from Santa Cruz) 1:100 – dilute 1uL Ab in 100uL block

anti-FLAG epitope (M2 from Kodak) 1:2500 – dilute 1uL Ab in 2500uL block

(done as 1 in 10, then 1 of that in 250)

9. You also need to make a humidified staining chamber. Take a 150mm petri dish, cut 2 paper towels in a circle to line the bottom. Wet the towels and pour off the excess water. Cut a circle of parafilm to place over the paper towels. Mark “top” across what will be the top of the dish to keep the orientation.

10. PRIMARY ANTIBODY: Using fine tweezers, move one coverslip face up to the humidified chamber. Pipet 50uL of the correct antibody solution on top of the coverslip. Do only one at a time, the coverslips must not dry out between solutions. Make sure to maintain the order of the coverslips so you know which are which. Don’t put the coverslips too close together – if the water on top of them touches, they will slip on top of one another. Incubate at room temp for 60 minutes.

11. WASH: Use the aspirator to suck off the primary antibody solution one CS at a time, and replace with 100uL PBS. Let the cells sit for 5 minutes, then repeat 2 more times (3 washes total).

12. SECONDARY ANTIBODY: When the cells are in the last wash, mix the secondary antibody dilutions. Secondary antibodies must be matched to the species of the primary antibody. Most primaries are either mouse or rabbit. Make sure you know which type of primary antibody you have for each antibody.

myc and FLAG antibodies are both mouse.

Secondary antibodies also come in colors, mostly red (Cy3, rhodamine, and texas red) or green (fluorescein, Alexa, Cy2). The microscope can also see blue (AMCA) but we rarely use this color. If you are staining for more than one protein each must be in a separate color. If you have cells expressing GFP, that is in the green channel so anything else you stain must be red.

We’ll use Cy3 anti-mouse (red).

We generally use the secondary antibodies at 1:500 in blocking solution.

Remove the last wash and add 50uL of the secondary antibody dilution to each CS. Incubate 60 minutes at room temp IN THE DARK ( to keep the Ab from fading).

13. WASH: wash 3 times with PBS for 5 minutes. One of the washes can contain Hoachst dye which will stain the nuclei blue. This allow you to see all the cells (expressing and not) in the fluorescent channel.

14. MOUNT: Place a drop of “Aquamount” on a glass slide for each coverslip. Don’t do more than one slide at a time as the mounting solution may dry before you are ready. You can put up to 6 small coverslips on a single slide. Make sure to label the slides on the white part, and keep track of the order of the coverslips on the slide in your notebook. Lift the CS with your fine tweezers, and suck off the excess fluid. Place the coverslip on the aquamount drop cell slide DOWN (flipped over from how you did the staining). Place the cells back in the dark to dry, then view on the fluoescent scope. The slides can be stored in the dark at 4 degrees for several months to a year.