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We recommend that you first download and read the PDF file containing the step-by-step protocol and solution recipes. Using that as your guide, you can then follow the protocol below and play Quicktime movies demonstrating key steps. We have also included higher resolution stills which provide close-ups of certain steps of the protocol, and you can view those by clicking on their thumbnail images.

Special Feature: Complete immunostaining protocol in one Quicktime movie

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The movie stars Dr. Archa Fox.

Protocol

Starting material: we grow our cells on 18 x 18 mm square or 19 mm circular coverslips in tissue culture dishes

Step 1-2:
Rinse cells with PBS (optional step) and fix as desired. If you're testing a new antibody, it's worth trying more than one fixation type/time. We generally use two different types of paraformaldehyde fixation: 1. warm 3.7% PFA in PHEM buffer for 10 min (good for preserving microtubules in mitotic cells) and 2. room temperature 3.7% PFA in CSK buffer (5 min only for staining speckles/paraspeckles/other splicing factors, can go to 10 min for most other antigens). We also use a methanol fixation for certain antibodies. Once fixed, rinse cells well with PBS.

Click here to view Quicktime movie showing the fixation and rinse steps (1.3 Mb).

Step 3-4:
Permeabilize with 1% Triton X-100 in PBS for 10 minutes at room temperature and rinse well with PBS.

Steps 5-6:
Set up humidified chamber (box with a lid and water-saturated tissue in it; bottom lined with parafilm) and place coverslips as desired (e.g. to use low solution volumes, in the range of 20-40 ul, pipette antibody directly onto parafilm and carefully place coverslip upside down, i.e. with cells facing down, on top of the drop of solution; if you're using solution volumes greater than 50 ul, you can place the coverslip on the parafilm with the cells facing up and pipette the antibody directly onto it).

Before adding antibody, however, block non-specific antibody binding sites by incubating cells for 10 min in blocking buffer (PBS + 0.1% Tween + 1% serum). After this step, incubate cells with primary antibodies diluted in blocking buffer for 35 mins to 1 hour. If you have never used the antibody before, try several dilutions (low of 1:100 to high of 1:1000, for example).

Step 7:
Transfer coverslips to 6-well plates for PBS washes (3 x 10 min). When washes are done, transfer back to humidified chamber for secondary antibody incubation). Incubate coverslips for 35 mins to 1 hour with the appropriate secondary antibodies diluted in blocking buffer, and wash 3 X 10 minutes with PBS. We use fluorophore-conjugated secondary antibodies from Jackson Immunochemicals. Transfer back to the 6-well plate for final PBS washes (3 x 10 min), and then stain with DAPI if desired (1:15,000 dilution in water of a 5 mg/ml stock). A good RNA stain, which fluoresces in the red channel, is Pyronin Y (1:10,000 dilution in water of a 10 mM stock; must stain with DAPI first). Wash well with PBS.

Step 8:
Mount the cells for immunofluorescence analysis. We commonly use either MOWIOL/Dabco, which hardens, or the liquid mounting media Vectashield (Vector Labs), in which case the coverslips must be sealed with nail varnish). Slides can be stored in the fridge or freezer until analyzed by microscopy.