Immunofluorescence Staining

Immunofluorescence is a technique allowing the visualization of a specific protein or antigen in cells or tissue sections by binding a specific antibody chemically conjugated with a fluorescent dye such as fluorescein isothiocyanate (FITC). There are two major types of immunofluorescence staining methods: 1) direct immunofluorescence staining in which the primary antibody is labeled with fluorescence dye, and 2) indirect immunofluorescence staining in which a secondary antibody labeled with fluorochrome is used to recognize a primary antibody. Immunofluorescence staining can be performed on cells fixed on slides and tissue sections and examined under a fluorescence microscope or confocal microscope.

The method relies on proper fixation of cells to retain cellular distribution of antigen and to preserve cellular morphology. After fixation, the cells are exposed to primary antibody directed against the protein of interest, in the prescence of permeabilizing reagents to ensure antibody access to the epitope. Following incubation with the primary antibody, the unbound primary antibody is removed and the bound primary antibody is then labeled by incubation with a fluorescently tagged secondary antibody directed against the primary antibody host species.


1XPhosphate Buffered Saline (PBS)
4% Formaldehyde: methanol free, use fresh and store opened vials at 4°C in dark. Dilute with 1X PBS to make a 4% formaldehyde solution.
0.1%Triton X-100 (prepared with 1× PBS)
Blocking Buffer: 1X PBS/5% normal serum (Sharing the same or similar species with secondary antibodies)
Antibody Dilution Buffer: To prepare 10 ml, add 30 µl Triton X-100 to 10 ml 1X PBS. Mix well then add 0.1g BSA.
Mounting medium: 50% glycerol with 0.1%(w/v) p-phenylenediamine in PBS or use Fluormount G

A. Sample Preparation

Cultured Cell Lines (IF-IC)
NOTE: Cells should be grown, treated, fixed and stained directly in multi-well plates, chamber slides or on coverslips.

1. Aspirate liquid, then cover cells to a depth of 2–3 mm with 4% formaldehyde diluted in warm PBS.
NOTE: Formaldehyde is toxic, use only in a fume hood.

2. Allow cells to fix for 15 min at room temperature.

3. Aspirate fixative, rinse three times in 1X PBS for 5 min each.

4. Permeate the cells at room temperature for 15 minutes with 0.1%Triton X-100 (prepared with 1× PBS)

5. Wash the cover glass with 1× PBS for 5 minutes and repeat 3 times.
Proceed with Immunostaining (Section B).

Frozen/Cryostat Sections (IF-F)

1. For fixed frozen tissue proceed with Immunostaining (Section B).

2. For fresh, unfixed frozen tissue, fix immediately, as follows:

a. Cover sections with 4% formaldehyde diluted in warm 1X PBS.
b. Allow sections to fix for 15 min at room temperature.
c. Rinse slides three times in PBS for 5 min each.
d. Permeate the cells at room temperature for 15 minutes with 0.1%Triton X-100 (prepared with 1× PBS)
e. Rinse slides three times in PBS for 5 min each.
Proceed with Immunostaining (Section B).

B. Immunostaining

NOTE: All subsequent incubations should be carried out at room temperature unless otherwise noted in a humid light-tight box or covered dish/plate to prevent drying and fluorochrome fading.

1. Block specimen in blocking buffer for 30 min.
2. Aspirate blocking solution, apply diluted primary antibody.
3. Incubate overnight at 4°C.
4. Rinse three times in 1X PBS for 5 min each.
NOTE: If using a fluorochrome-conjugated primary antibody, then skip to Step 7.

5. Incubate secondary antibody diluted in antibody dilution buffer for 1–2 hr at room temperature in the dark.
6. Rinse three times in 1X PBS for 5 min each.
7. Invert Coverslip slides cell-side-down, on one drop mounting medium. Gently blot with paper towel, then seal edge by painting with nail polish. Let dry.
8. For best results, allow mountant to cure overnight at room temperature. For long-term storage, store slides flat at 4°C protected from light (6-12 month).


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