Immunocytochemistry and immunofluorescence staining protocol

Find out how to use fluorescent antibodies or dyes to detect target antigens within cells (6:59 minutes). ​

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Slide preparation

View our Counting cells using a hemocytometer protocol here if you need more detailed infomation.

1. Coat coverslips with polyethylineimine or poly-L-lysine for 1 h at room temperature.
2. Rinse coverslips well with sterile H₂O (three times 1 h each).
3. Allow coverslips to dry completely and sterilize them under UV light for at least 4 h.
4. Grow cells on glass coverslips or prepare cytospin or smear preparation.
5. Rinse briefly in phosphate-buffered saline (PBS).
   
   For wash buffer we recommend 1x PBS 0.1% Tween 20.
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Fixation

The cells may be fixed using one of two methods:

1. Incubating the cells in 100% methanol (chilled at -20°C) at room temperature for 5 min.
2. Using 4% paraformaldehyde in PBS pH 7.4 for 10 min at room temperature.

The cells should be washed three times with ice-cold PBS.

Antigen retrieval (optional step)

Certain antibodies work best when cells are heated in antigen retrieval buffer. Check the product information for recommendations for each primary antibody being used.

1. Preheat the antigen retrieval buffer (100 mM Tris, 5% [w/v] urea, pH 9.5) to 95°C. This can be done by heating the buffer in a coverglass staining jar which is placed in a water bath at 95°C.
2. Using a small pair of broad-tipped forceps, place the coverslips carefully in the antigen retrieval buffer in the cover glass staining jar, making note of which side of the coverslips the cells are on.
3. Heat the coverslips at 95°C for 10 min.
4. Remove the coverslips from the antigen retrieval buffer and immerse them, with the side containing the cells facing up, in PBS, in the 6-well tissue culture plates.
5. Wash cells in PBS three times for 5 min.
   
   

Permeabilization

If the target protein is intracellular, it is very important to permeabilize the cells. Methanol fixed samples do not require permeabilization.

1. Incubate the samples for 10 min with PBS containing either 0.1–0.25% Triton X-100 (or 100 μM digitonin or 0.5% saponin). Triton X-100 is the most popular detergent for improving the penetration of the antibody. However, it is not appropriate for membrane-associated antigens since it destroys membranes.
2. The optimal percentage of Triton X-100 should be determined for each protein of interest.
3. Wash cells in PBS three times for 5 min.
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Blocking and immunostaining

1. Incubate cells with 1% BSA, 22.52 mg/mL glycine in PBST (PBS+ 0.1% Tween 20) for 30 min to block unspecific binding of the antibodies (alternative blocking solutions are 1% gelatin or 10% serum from the species the secondary antibody was raised in (typically goat serum, or donkey serum): see antibody datasheet for recommendations).
2. Incubate cells in the diluted antibody in 1% BSA in PBST in a humidified chamber for 1 h at room temperature or overnight at 4°C.
3. Decant the solution and wash the cells three times in PBS, 5 min each wash.
4. Incubate cells with the secondary antibody in 1% BSA for 1 h at room temperature in the dark.
5. Decant the secondary antibody solution and wash three times with PBS for 5 min each in the dark.
   
   

Multicolor staining (optional step)

To examine the co-distribution of two (or more) different antigens in the same sample, use a double immunofluorescence procedure. This can be performed either simultaneously (in a mixture) or sequentially (one antigen after another).

Ensure you have antibodies for different species and their corresponding secondary antibodies. For example, rabbit antibody against antigen A, mouse antibody against antigen B. Alternatively, you can use directly conjugated primary antibodies conjugated to different fluorophores.

Simultaneous incubation

1. Incubate cells with blocking solution for 30 min.
2. Incubate cells with both primary antibodies in 1% BSA in PBST in a humidified chamber for 1 h at room temperature or overnight at 4°C.
3. Decant the solution and wash the cells three times in PBS, 5 min each wash.
4. Incubate cells with both secondary antibodies in 1% BSA for 1 h at room temperature in the dark.
5. Decant the secondary antibody solution and wash three times with PBS for 5 mins each in the dark.

Sequential incubation

1. First blocking step: incubate cells with the first blocking solution (10% serum from the species that the secondary antibody was raised in) for 30 min at room temperature.
2. Incubate cells with the first primary antibody in 1% BSA or 1% serum in PBST in a humidified chamber for 1 h at room temperature or overnight at 4°C, 1% gelatin or 1% BSA.
3. Decant the first primary antibody solution and wash the cells three times in PBS, 5 min each wash.
4. Incubate cells with first secondary antibody in 1% BSA in PBST for 1 h at room temperature in the dark.
5. Decant the first secondary antibody solution and wash three times with PBS for 5 min each in the dark.
6. Second blocking step: incubate cells with the second serum, (10% serum from the species that the secondary antibody was raised in) for 30 min at room temperature in the dark.
7. Incubate cells with the second primary antibody in 1% BSA or 1% serum in PBST in a humidified chamber in the dark for 1 h at room temperature, or overnight at 4°C.
8. Decant the second primary antibody solution and wash the cells three times in PBS, 5 min each wash in the dark.
9. Incubate cells with second secondary antibody in 1% BSA for 1 h at room temperature in the dark.
   
   Decant the second secondary antibody solution and wash three times with PBS for 5 min each in the dark.
   
   

If you have to detect more than two antigens, continue following steps 1–5 for the rest of the antibodies.

Counter staining

1. Incubate cells on 0.1-1 μg/mL Hoechst or DAPI (DNA stain) for 1 min.
2. Rinse with PBS.
   
   

Mounting

1. Mount coverslip with a drop of mounting medium.
2. Seal coverslip with nail polish to prevent drying and movement under microscope.
3. Store in dark at -20°C or +4°C.
   
   

• View our other ICC/IF resources
• Get images like the one below with  Alexa Fluor®  conjugated  antibodies

A. Annexin V labeled with Alexa Fluor^® 488 in frozen rat placenta section by IHC (immunohistochemistry). Green - Anti-Annexin V antibody [EPR3980] (Alexa Fluor^® 488) (ab201540); Red - Anti-Tubulin antibody [YOL1/34] (Alexa Fluor^® 647) (ab195884); Blue - DAPI.

B. Lamin B1 labeled with Alexa Fluor^® 488 in HUVEC cells using Alexa Fluor^® 488-conjugated secondary antibody.  Green - Anti-Lamin B1 antibody (ab16048), secondary antibody Goat Anti-Rabbit IgG H&L (Alexa Fluor^® 488) (ab150077); Red - Anti-alpha Tubulin antibody (ab7291), secondary antibody  Goat anti-mouse IgG H&L (Alexa Fluor^® 647) (ab150115);  Blue - DAPI.

See advantages of Alexa Fluor®  secondary antibodies for ICC/IF

Alexa Fluor^® is a registered trademark of Life Technologies. Alexa Fluor^® dye conjugates contain(s) technology licensed to Abcam by Life Technologies.

Webinar transcript

Immunocytochemistry or ICC is a technique that uses antibodies to visualize the localization of particular proteins within single cells. This video outlines slide preparation from suspension cells followed by fixation, permeabilization, blocking, and antibody incubation. Assemble the cytospin equipment before removing the cells from the incubator.

The cytospin deposits thin layer preparations of suspension cells onto slides. Load the cells and set it running. The cell density, loading volume, and spin speed should be optimized for each cell type. For example, larger cells require a slower speed. Unload the slides carefully and place them in a slide tray to dry ... before transferring them onto a slide rack.

Ideally, the cell should be fixed immediately. Therefore, place the rack of slides into a pot of fixative and incubate for 5-10 minutes. Next, wash the slides by transferring them sequentially into two pots of Phosphate Buffered Saline, PBS.

Once fixed they can be stored for a short period of time in the fridge but keep them moist. The type of fixative used, and the fixation time will require optimization. The most commonly used fixatives are paraformaldehyde and methanol. However, it is advisable to check the datasheets of the primary antibodies you intend to use for guidance.

Remove the slides from the rack and tap off excess PBS. Draw a hydrophobic barrier around the cells using a PAP pen. This will keep reagents pooled in a small droplet over the cells. It is important to ensure the cells do not dry out during this process. Carefully pipette the permeabilization buffer onto the cells and incubate for five minutes.

The permeabilization step allows antibodies to access intracellular epitopes and will need optimizing depending on the cell type, antigen, and intended antibody.

There are a number of different permeabilization agents that can be used. For example, Triton X-100 and for very gentle permeabilization, Tween-20. You do not need to permeabilize if the cells were previously fixed in methanol.

Once the cells are permeabilized, tap excess buffer off the slides and wash in Phosphate Buffered Saline plus Tween-20, PBST, on a shaker for five minutes. Repeat with fresh PBST for a total of three wash steps. Once the washes are complete leave the slides in your wash buffer to avoid them drying out.

While your slides are washing, dampen some paper towel and use it to line a slide tray. This will help to keep your slides moist during the next step. Remove excess buffer from your slides and re-draw your PAP pen circles as they may have partially washed off.

To prevent non-specific antibody binding, the cells need to be blocked. In this case, the cells are blocked in 10% serum and one percent Bovine Serum Albumin, BSA, for one hour at room temperature. The type of blocking buffer and incubation time should be optimized for your experiment.

Note; if the cells have been fixed in paraformaldehyde it is common practice to wash with glycine before blocking to quench any remaining fixative. Ideally, the species of serum in the blocking buffer should match the species the secondary antibody was raised in to avoid any cross-reactivity.

Whilst the cells are in blocking buffer, prepare the primary antibodies. After blocking, the cells need to be washed three times for five minutes in PBST as before. Remove any excess wash buffer and re-draw the PAP pen circles as they may have partially washed off.

Place your slides in the line slide tray and add your primary antibody solution. Do this one slide at a time to avoid drying. Place a lid on your tray and carefully transfer it to a fridge to incubate overnight.

Wash the slides three times for five minutes in PBST to remove any unbound primary antibody. If the primary antibodies weren't directly conjugated, incubate the cells for one hour at room temperature with secondary antibodies. Nuclear stains such as Hoechst can be included at this point too. Then wash the slides three times again in PBST.

Tap excess buffer off your first slide and place on a slide tray. It is best to deal with one slide at a time to avoid drying. Add a droplet of compatible mounting medium onto the cells ensuring no air bubbles are created.

Using a pair of forceps lower the coverslip down onto the mounting medium allowing the liquid to spread out gradually. Avoid putting any pressure on your slides as this may damage your cells. The slides are now ready for imaging using a confocal microscope or can be stored in the fridge for a short period of time.