Center for Bio-Image Informatics

Immunocytochemistry for confocal microscopy

Procedure
Immunocytochemistry for confocal microscopy

1. Agarose Embedding (Mouse Tissue)

1. Take eye piece (i.e. half-eye cup) rinse in cold (4°C) 1XPBS (pH = 7.4) if the tissue was stored in 4% Paraformaldehyde. 15min rinses 1hr. rinse.

2. Melt the Agarose: Place solid agarose (in tube) in a beaker full of water, with the tube lid loose. Put the beaker in the microwave for 2 min (or until melted).

3. Insert a thermometer directly into the melted agarose and transfer agarose tube to a second beaker filled with room temperature water. Stir agarose.

   Allow the agarose to cool to 44-45°C (any lower and the agarose will harden and any hotter the tissue will autoflourecence).

4. Embed the tissue in an agarose block by pouring agarose into a small weigh boat. Then transfer the tissue from PBS to liquid agarose and orient it (as shown on the next page in final form) using a spatula. (Note: Ideally the tissue should be put in liquid agarose at 40-42°C. If you pour at 44-45°C, a few seconds of cooling in the boat is all that is needed.) When the tissue is on the spatula use a Kim wipe and CAREFULLY wipe around the tissue. When you first put the tissue in agarose, watch it for about 1-2min to make sure the tissue does not move. If it does, reposition it quickly with a spatula.

5. Place the tissue-agarose block in a Petri dish on a wet paper towel making sure there is ample water for the block to keep hydrated. Cover the dish with plastic wrap and store at 4°C for 30 min to 1-hour minimum to harden (can be kept overnight or longer as long as the block is well hydrated).

2 Sectioning (Embedded Mouse Tissue

1. Break a double edged razor blade in half, clean with 70% Ethanol, and mount in vibratome.

2. Cut small block around tissue in agarose. Tissue should be oriented as shown below with the longest side vertically and the sides of the block should be sloped (pyramidal).

3. Glue the block on to the vibratome chuck with Krazy Glue.

4. Put the chuck (with tissue glued) and some PBS into the vibratome well until the block is entirely submerged in cold (4°C) 1XPBS. Adjust blade so tha t it cuts entirely through the block.

5. Cut sections at 100 µm thickness and transfer them to cold PBS using a camel hair paint brush. Do not keep the first section cut (it will be of unknown thickness).

6. Rinse once with 1X PBS.

7. Place sections in solution (Block) for immediate use (see immuno run protocol) or 4% paraformaldehyde fix for short or long-term storage.

3. Immuno Run

1. Rinse: If sections were fixed in 4% paraformaldehyde then rinse in cold (4ºC) 1X PBS. 3 15min rinses 1hr. rinse. In a standard run 2-6 sections are used per well, where each well will be for a particular primary antibody (or antibody combination).

2. Block: Make 1:20 NDS (Normal Donkey Serum) in cold 1X PBTA (i.e. 25 µl NDS in 475 µl 1X PBTA) and add 500 µl for each cup/well. Incubate 2 hrs to overnight (or longer) at 4ºC

3. Primary Antibodies: Make all antibody dilutions in cold (4ºC) 1X PBTA. Make 500 µl total solution for each cup/well. Remove Blocking solution and add primary antibody solution. Incubate overnight at 4ºC.

4. Rinse Remove (and save) primary antibodies (store most at 4ºC). Rinse with cold (4ºC) 1X PBTA. 3 15min rinses 1hr. rinse.

5. Secondary Antibodies: Dilute secondary antibodies (i.e. DAM-Cy3) to 1:200 in cold (4ºC) 1X PBTA. Strepavadin antibodies are diluted 1:100. Remove fina l rise and add diluted secondary antibodies. Incubate overnight 4ºC.

6. Rinse: Remove (and discard) secondary antibodies from cups/wells and 3 15min rinses 1 1hr. rinse.

7. Mounting on Slides: Lift sections GENTLY out of well with a spatula. Cut the section (if necessary) while on the spatula. If section dries on spatula, don't rip it off, instead use PBTA and rinse section carefully.

   Push the section off the spatula and on to a coverslip (thickness=0 mm) using a razor blade (blunt side works well). Place up to 4 sections per coverslip

   Wick away excess PBTA with a Kimwipe and place a drop of n-propyl-gallate on the coverslip (use a pastuer pipette).

   Invert coverslip onto a glass slide. Lower slide until it briefly touches n-propyl-gallate. Allow cover slip to "suck-up" onto the slide and slowly invert. If bubbles cover any of the tissue, the cover slip can be lifted with a razor and spatula (but be slow and careful). Wick away excess or ad d a small amount (if not enough) of n-propyl-gallate at edge of cover slip.

   Seal cover slip with nail polish.