Immunofluorescence General Protocol
The protocols described below are for general application. Any product specific protocol supercedes these general recommendations.
II. Indirect Immunofluorescence (tissues) Protocol
III. Indirect Immunofluorescence (cells) Protocol
Immunofluorescence is a technique that allows 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. Immunofluorescence stained samples are examined under a fluorescence microscope or confocal microscope.
Because fluorescent dyes such as fluorescein and rhodamine can be coupled to antibodies without destroying their specificity, the conjugates can complex with antigen and be visualized via fluorescence microscopy. The microscope excites the chosen dyes by light of one or more wavelengths, which in turn emits light at a characteristic wavelength captured by a selective filter. The image is then projected with localized regions of fluorescence indicating different antigens labeled by antibodies of distinctive color.
|Common Fluorescent Tags||Excitation (nm)||Emission (nm)|
The antibody is itself conjugated with the fluorochrome and applied directly to a monolayer of cells or to frozen tissue on a slide. When examined with a fluorescence microscope, the antibody labeled with the fluorescent compound identifies the localized antigen.
Unlike direct immunofluorescence, indirect immunofluoresence is a double-layer technique. The unlabeled antibody is applied directly to the tissue substrate and then treated with a fluorochrome-conjugated anti-IgG. There are several advantages to this technique, and it is typically used more frequently than the direct method. Because several fluorescent anti-immunoglobulins can bind to each antibody present in the first layer, this produces brighter fluorescence than in the direct method. It is also more time-efficient since there is only one fluorescent-labeled reagent, the anti-IgG prepared during the lengthy conjugation process. A general protocol for only the indirect immunofluorescence technique is presented here.
II. Immunofluorescence (tissues) Protocol
A. Materials required
Slide racks & tray
Staining dishes with lids
PAP pen & Transfer pipettes
Deionized water (ddH2O)
PBS (Phosphate Buffered Saline)
Fluorescent-labeled secondary antibody
B. Sample preparation
1. Samples are perfused or dissected and fixed in 4% paraformaldehyde fixative.
2. Transfer the tissue to 20% sucrose in PBS, leave overnight at 4°C.
3. Transfer the tissue to 30% sucrose in PBS, leave at 4°C to impregnate fully. When the tissue sinks, it is fully impregnated.
4. Freeze and cut 5-20mm thick crysostat sections
C. Blocking Step
5. Incubate slides for 1 hour in 5% normal serum (serum of secondary antibody host) in PBS.
6. Wash with PBS, 3x 5 minutes.
D. Primary antibody incubation step
7. Dilute primary antibody in PBS, 0.1% normal serum.
8. Incubate sample with primary antibody for 1 hour at RT or overnight at 4°C.
9. Wash 3x with PBS.
E. Secondary antibody incubation
10. Incubate with fluorescence-conjugated secondary antibody in PBS, 0.1% normal serum for 1-2 hours at RT in the dark. (From this point on, keep slides in the dark.)
11. Wash 3x with PBS. To reduce background can include 0.1% Tween 20 in PBS.
F. Fluorescence detection
13. Examine using fluorescence microscopy immediately or store flat at 4°C in the dark.
III. Immunofluorescence (cells) Protocol
A. Materials required
Distilled water (dH2O)
PBST: 1X PBS, 0.1% Triton X-100. To prepare 1L, add 100ml 10X PBS to 900ml dH2O. Add 1ml Triton X-100 and mix.
Fluorochrome-conjugated secondary antibody
B. Specimen Preparation
Note: Cells should be grown, treated, fixed, and stained directly in multiwell plates, chamber slides, or on coverslips.
1 To increase cell adherence, treat coverslips with a 1:10 dilution of poly-lysine solution at room temperature for 5 minutes.
2. Plate cells at appropriate dilution and grow until cells reach desired confluence (~70%)
3. Aspirate media
4. Rinse cells briefly in PBS.
C. Fixation step
5. Aspirate PBS, cover cells with 2-4% formaldehyde in PBS (work in fume hood). Allow cells to fix for 15 minutes at RT.
6. Aspirate fixative, rinse 3x in PBS for 5 minutes each.
D. Permeabilization (optional)
Methanol and acetone Fixation result in permeabilized cell preparations. For paraformaldehyde fixed preparations, treat with 0.2% Triton X-100 for 5 min or alternatively with -20°C methanol for 5 minutes.
7. Methanol Permeabilization Step: After formaldehyde fixation, cover cells with ice-cold 100% methanol (use enough to cover cells completely to a depth of 3-5mm, do not let cells dry), incubate cells in methanol for 10 minutes in freezer.
8. Rinse in PBS for 5 minutes.
E. Blocking step
Note: All incubations should be carried out at RT unless otherwise noted in a humid light-tight box or covered dish to prevent drying and to prevent exposure of fluorochrome to light.
9. Block sample in 5% normal serum from same species as secondary antibody (eg. normal goat serum, normal donkey serum) in PBS,1%Triton X-100 for 1 hour. (Recommended Blocking Buffers can vary between 1-2% Bovine Serum Albumin, Fetal Bovine Serum Albumin, or 2% Non-fat dry milk in TBS (PBS) with or without 0.2% Tween 20 and 0.02% Sodium Azide.)
F. Primary antibody incubation
10. Dilute primary antibody in PBS, 0.1% Triton. Typical volumes are: 50-100ul per section, 25-50ul per coverslip, chamber, or well (48 or 96 well plate).
11. Aspirate blocking solution, apply diluted primary antibody.
12. Incubate overnight at 4°C with gentle agitation or rocking.
13. Rinse 3x in PBS, 0.1% Triton X-100 for 5 minutes each.
G. Secondary antibody incubation
14. Incubate with fluorochrome-conjugated secondary antibody diluted in PBS, 0.1% Triton X-100 for 1-2 hours at RT in dark. (From this point on, slides need to be kept in the dark.)
15. Rinse in PBS, 0.1% Triton X-100.
H. Fluorescence detection
16. Mount coverslips.
17. Examine using fluorescence microscopy immediately or store flat at 4°C in the dark.
IV. Troubleshooting for Immunofluorescence
Problem: Background fluorescence
1. Dilute the fluorochrome-conjugated antibody further. If it is too concentrated, background may increase due to an increase in non-specific interactions.
2. Be sure to include a blocking step to reduce background by blocking non-specific interactions between the primary antibody and the cell surface or intracellular structures.
3. Use F(ab')2 fragments where background may be due to binding of the whole molecules of primary or secondary antibodies to Fc-receptors. F(ab')2 fragments of most secondary antibodies are readily available.
4. Incubate fresh tissues or cells with normal serum in the presence of sodium azide prior to addition of the primary antibody.
5. Use cross-adsorbed secondary antibodies to remove unwanted cross-reactivities between labeled antibodies and any other immunoglobulins inherent or added to the experimental system.
6.. Employ direct immunofluorescence. Indirect labeling can boost overall signal intensity, but the use of secondary antibodies can result in extensive non-specific binding, compromising signal to background ratio.
7. For fixed tissue, remove free aldehyde groups with 0.1% sodium borohydride in PBS for 30 mins prior to staining.