Immunofluorescence staining protocol

1. Cell preparation
   • Adherent cells:
     • Grow desired cells on glass coverslips (preferably thickness #1) in six-, 12- or 24-well plates.
       Note: For analysis such as live imaging, grow cells on round glass coverslips.
   • Suspension cells:
     • Coat coverslips with excess polylysine (0.01% solution) for 10 minutes at room temperature. Aspirate the excess and let the coverslips dry. Rinse coverslips with water and place them in six-, 12- or 24-well plates.
     • Wash cells with PBS and resuspend. Add the resuspended cells onto the polylysine-coated coverslips and let them settle for 30–60 minutes. Aspirate the excess followed by a gentle rinse with PBS.
       
       
2. Fixation
   For adherent cells, wash the coverslips with PBS at least three times to completely remove the media.
   
   Replace PBS with one of these fixation solutions below.
   
   There are different methods for fixing cells:
   • 4% paraformaldehyde (PFA) in PBS for 30 minutes or more at room temperature
     Note: This is a common method used for cell fixation. The 4% PFA can be made by diluting 16% PFA stock. It is recommended to prepare 4% PFA fresh.
   • 100% chilled methanol at -20°C for 10 minutes
     Note: This method fixes cells rapidly, thus useful for cytoskeleton staining, but can cause shrinkage of cells. If cells are fixed by this method, the permeabilization step can be skipped. In addition, this method cannot be used for fluorescent conjugated proteins (e.g. GFP), transfected cells and phalloidin staining.
   
   After fixation, wash the coverslips with PBS at least three times to remove the fixation buffer.
   
   
3. Permeabilization
   • Permeabilize with 0.1–0.05% Triton X-100 in PBS (containing 100 mM glycine) for 20 minutes at room temperature.
     Note: Permeabilization is used when staining for internal antigens.
   • After permeabilization rinse the coverslips with PBS at least three times.
     
     
4. Blocking
   There are different blocking solutions that can be used. Please check with the primary antibody information sheet for specifications.
   • Block with 2–5% animal serum, BSA or milk diluted in PBS for one hour at room temperature.
     
     
5. Incubating with primary antibody
   • Incubate the coverslips with diluted primary antibody in 1% blocking buffer for one hour at room temperature.
     Note: In general primary antibodies are diluted at (1:100–1:1,000). For specifications on dilution please check with the antibody’s information sheet. In order to conserve antibodies, the coverslips can be picked up by a tweezer and flipped onto a drop of antibody solution (75–100 μL) that can be placed on a piece of parafilm.
   • Rinse the coverslips with PBS at least three times.
     
     
6. Incubating with secondary antibody
   • Incubate the coverslips with diluted secondary antibody in 1% blocking buffer for one hour at room temperature.
     Note: In general, secondary antibodies are diluted at (1:1,000-1:1,500). Secondary antibodies from Jackson are the preferred choice. It is recommended to cover the plates with foil or place them in dark in order to limit bleaching of samples from this stage of the procedure.
   • Rinse the coverslips with PBS at least three times.
     Note: If cells will be stained with DAPI, the coverslips need to be rinsed with water at this stage before proceeding with DAPI staining. In general, for DAPI staining, incubate the coverslips with 1:10,000 diluted DAPI for 10 minutes then rinse with water.
     
     
7. Mounting
   There are different mounting media that can be used: DAKO, Mowiol or PBS:Glycerol (1:1) followed by clear nail polish to seal coverslips.
   • Flip the coverslip onto a drop of mounting media placed on a slide.
     Note: Remove any air bubbles trapped between the coverslip and slide.
     
     

   Leave the slides to dry overnight.