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Protocol: Immunofluorescence Staining of Cells for Microscopy

There are many variations on IF protocols, and steps may need to be optimized for different targets or applications. Some epitopes may require specific fixation conditions for detection. This is our basic protocol for staining adherent cells in dishes or cells grown on coverslips.

Materials required:

Workflow overview:
  1. Fix (≤20 min.) (optional stopping point)
  2. Block/permeabilize (30 min.) (optional stopping point)
  3. Primary antibody (2 hours or overnight)
  4. Washes (20-30 min.)
  5. Secondary antibody (30 min. to 2 hours)
  6. Washes, (20-30 min.)
  7. Mount (optional stopping point)
  8. Image


    1. Rinse cells twice with PBS or HBSS to remove cell culture medium. Use the same volume for washes as you would for cell culture medium (we use 100 uL per well of a 96-well plate). For some cell types, buffer with Ca2+/Mg2+may be necessary to prevent cell rounding and detachment. Prior to fixation, we prefer to use HBSS + Ca2+/Mg2+ for adherent cells.
    2. Fix cells. We usually use 4% paraformaldehyde/PBS, 20 min. at room temperature. Alternatively, fix cells in pre-chilled methanol at -20°C for 5-10 min.
      Note: Check the information provided by the primary antibody supplier to see if a specific fixation method is recommended. If the optimal fixation conditions are unknown, it may be necessary to test different fixation methods for a specific antibody or target epitope.
      Note: Methanol fixation is not compatible with phalloidin staining.
    3. Rinse three times with PBS to remove traces of fixative.
      Note: In our experience, cells can be stored in PBS after fixation for several weeks. Keep samples well-sealed or in a humidified box to avoid evaporation of buffer.
    4. Block and permeabilize cells in PBS + 2% fish gelatin + 0.1% Triton® X-100.
      Optional: You can store samples at 4°C for several weeks at this point. Keep samples well-sealed or in a humidified box to avoid evaporation of buffer.
      Note: When using some highly charged fluorescent dyes, specialized blocking buffers such as our TrueBlack® IF Background Suppressor System may reduce background.
    5. Dilute primary antibody in fresh blocking/permeabilization buffer at the concentration recommended by the antibody supplier.
      Note: You may need to perform a titration experiment to determine the optimal concentration of primary antibody.
    6. Add enough diluted antibody solution to cover cells completely. We usually use 50-100 uL per well of a 96-well plate.
      Note: For cells on coverslips, add 50-100 uL of diluted antibody solution and overlay with a piece of Parafilm® to spread solution evenly over the specimen, making sure there are no bubbles. Keep samples in a humidified chamber to avoid evaporation.
    7. Incubate 1-2 hours at room temperature or overnight at 4°C (in our experience, 4°C overnight gives the best results). If using fluorescently labeled primary antibodies, protect samples from light.
      Note: Other stains such as nuclear counterstains, lectins, or phalloidin conjugates can be added together with labeled antibodies at this step, or at step 10 if using labeled secondary antibodies.
    8. Rinse cells twice with PBS, then wash 3 x 5 min. with PBS.
      Note: Alternatively, rinse cells twice with PBS, incubate in PBS for 30 min., then rinse with PBS. Cells can be left in PBS for longer times without negatively affecting staining.
    9. If using directly labeled primary antibodies, proceed to step 12. If using labeled secondary antibodies, proceed to step 10.
    10. Dilute secondary antibody in blocking/permeabilization buffer at 1 ug/mL. Cover cells with secondary antibody solution as in step 5 and incubate for 30 min. to 2 hours at room temperature, protected from light.
    11. Wash cells as in step 8.
    12. Mount samples in fluorescence antifade mounting media such as EverBrite™ Mounting Medium (medium with DAPI can be used for blue nuclear counterstaining). For chambered coverglass or multi-well coverglass plates, remove all traces of buffer and add enough mounting medium to completely cover the cells.
      Note: For coverslips, wet-set or hard-set mounting medium may be used. Follow mounting medium instructions for mounting coverslips. If wet-set mounting medium is used, the edges of the coverslip must be sealed with nail polish or CoverGrip™ Coverslip Sealant (recommended) before imaging.
    13. Store samples in the dark at 4°C until ready to image. Samples can be stored in mounting medium at 4°C for six months or longer.
      Note: Phalloidin staining is less stable than antibody staining. Staining with most phalloidin conjugates is stable at 4°C for several days, but for best results it should be imaged within 24 hours.
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