Protocols: Immunofluroescence Technique

  1. Fix cells in 2% formaldehyde in PBS/pH 7.4 for 15 min. at 20oC. 2% formaldehyde is made up fresh prior to use by dissolving the appropriate amount of EM grade paraformaldehyde (Prill form, from Electron Microscopy Sciences) in PBS and heating on a hot plate in the hood (setting of 5-6) until the aldehyde goes into solution. Keep the bottle cap loosened so that pressure does not build up. Cool down to 20oC and pH to 7.4. Alternatively, cells may be fixed in 100% methanol at -20oC for 3 minutes. If methanol fixation is used skip to step 4.
  2. Wash in PBS 3 X 10 min.
  3. Permeabilize in 0.2% Triton X-100 plus 1% normal goat serum (NGS) in PBS/pH 7.3 for 5 minutes on ice.
  4. Wash in PBS + 1% NGS 3 X 10 min.
  5. Incubate in the appropriate concentration of primary antibody for 1 hour at room temp. in a humidified chamber. If using 22mm X 22mm square coverslips, 30 ul of diluted antibody is placed on the coverslip and the coverslip is inverted onto a glass slide. The slide is then placed in the humidified chamber which is incubated at room temp.
  6. Wash in PBS + 1% NGS 3 X 10 min.
  7. Incubate in secondary antibody (FITC or Texas Red conjugated) at a dilution of 1:50 for 1 hour in a humidified chamber at room temp.
  8. Wash in PBS 4 X 10 min.
  9. Mount coverslip with a drop of mounting medium (see Fluorescence Mounting Medium Protocol) and seal coverslip with clear nail polish to prevent drying and movement under the microscope.

From: Spector, D.L. and H.C. Smith. 1986. Exp. Cell Res. 163, 87-94.

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