Multicolor Immunofluorescence Staining Protocol

Related Links

Immunocytochemistry (ICC) Handbook

ICC Application Resources

Fluorochromes Selection for Multicolor ICC/IF

Sample Preparation for ICC/IF Experiments

Fixation & Permeabilization

Blocking for ICC/IF Assay

Antibody Selection in ICC/IF

Detection Methods for ICC/IF

Controls for ICC/IF experiments

Counterstaining & Mounting

Organelle Markers Guide

Secondary Antibody Handbook

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What is Multicolor Immunfluorescence Staining?

Double immunofluorescence staining, also known as or multicolor immunocytochemistry (ICC) or multicolor immunofluorescence (IF), is helpful to examine the distribution of two (or more) different antigens in the same sample of cells. Multicolor ICC/IF can be performed either simultaneously using an antibody cocktail or sequentially by probing one antigen after another. Simultaneous and sequential staining follow the same basic protocol, but considerable variation exists in the blocking and antibody incubation steps. In all multicolor staining protocols, the choice of either directly labeled primary or secondary antibodies is very critical. When picking fluorochrome colors for labeled antibodies, one must consider their fluorescent emission spectrum. Bleed through effects will be minimal if the selected fluorochromes have negligible spectral overlap. Also, it is recommended to use the dimmest fluorochrome to label the most abundant protein and vice versa. A negative control lacking primary antibody should always be included to check the specificity of staining. These precautions and the tips discussed in the protocols below will help to efficiently/accurately detect multiple proteins in immunofluorescence staining.

CELL CULTURE

1. Coat the coverslips with poly-L-lysine, dry and sterilize them.


Tip: Coverslips can be sterilized by dipping in ethanol and flaming them or by placing them into tissue culture dishes and exposing them to UV radiation (available in most tissue culture hoods).


2. Seed the cells on sterile glass coverslips (poly-L-lysine coated). Some cell types may require growth on other treated coverslips for proper adhesion. Grow the cells to semi-confluency.
   
   Tips: Multiple coverslips can be seeded in a single dish which helps reduce the number of dishes in the incubator. Keep all mechanical manipulations to a minimum to avoid compromising the quality of the final cell images. Always treat the cells and coverslips gently, never let them dry out and avoid dropping solutions directly on the cells.

FIXATION

1. Aspirate the culture medium from the dish or remove each coverslip as required with tweezers, and gently wash them with PBS at room temperature.


2. Incubate the coverslips in freshly prepared 4% paraformaldehyde – neutral PBS at room temperature for 10 minutes. Alternatively, the cells can be fixed for 10 minutes in chilled methanol (pre-equilibrated at -20°C) on ice.


Tip: Alternative fixation methods may be tested and compared to determine which is best at preserving the structure and epitope of the protein of interest.


3. Wash the coverslips of fixative in PBS for 2 minutes.

PERMEABILIZATION

1. Incubate the coverslips in 0.5% Triton X-100 in PBS at room temperature for five minutes. Test different detergents (ex. digitonin, Tween-20) in a range of concentrations to find the optimal condition that best preserves cell structure and the target protein.


Tip: A permeabilization step is not required with methanol fixation because Methanol acts as a fixative as well as cell permeabilization agent.


2. Wash the coverslips of the permeablization buffer by incubating in PBS for 5 minutes.

BLOCKING

1. IgG from the secondary antibody may bind non-specifically to the sticky sites on the cells which often leads to non-specific background signal. To avoid this issue, block the coverslips in 1-5% normal serum prepared in PBS for one hour at room temperature.


Tip: The normal serum block should be of the same species in which the secondary antibody has been raised. For example, if you are using a goat-anti rabbit secondary, then block with 5% normal goat serum.


2. Alternative blocking agents are 1% gelatin or 1-5% BSA in PBS.

MULTICOLOR IMMUNOFLUORESCENCE STAINING

Multicolor labeling experiments are best carried out by sequentially incubating cells with primary and secondary antibodies, however it may be performed by employing one of the following three options:

Option #1: Sequential incubations with unlabeled antibodies
This method is useful when the primary antibodies involved are from different hosts (e.g. mouse monoclonal against antigen-X, rabbit polyclonal against antigen Y, and sheep polyclonal against antigen Z ), and the antibodies display aggregates formation in the simultaneous incubation method.

1. Blocking step #1: incubate cells with blocking buffer solution #1 for 30 minutes at room temperature. This buffer solution contains 5% serum from the same species as the host of the labeled secondary antibody #1 (against primary antibody #1).


2. Primary incubation #1: incubate cells with the primary antibody #1 in 1% BSA or 1% serum in a humidified chamber for one hour at room temperature or overnight at 4°C.


3. Decant the antibody solution and wash the cells three times in PBS-T (5 minutes for each wash).


4. Secondary incubation #1 - Incubate cells with secondary #1 in 1% BSA for one hour at room temperature in the dark.


5. Decant the antibody solution and wash the cells three times in PBS-T (5 minutes for each wash).


Tip: Process the cells for next step immediately and do not let the cells dry out at any stage.


6. Blocking step #2: incubate cells with the blocking buffer #2 (5% serum from the species which is the same as the host of the secondary #2) for 30 minutes at room temperature in the dark.


7. Primary incubation #2: Incubate cells with primary antibody #2 in 1% BSA or 1% serum in a humidified chamber in the dark for one hour at room temperature, or overnight at 4°C.


8. Decant the antibody solution and wash the cells three times in PBS-T (5 minutes for each wash).


9. Secondary incubation #2: Incubate cells with secondary antibody #2 in 1% BSA for one hour at room temperature in the dark.


10. Decant the secondary antibody solution and wash three times with PBS for five minutes each in the dark.


Tip: Repeat steps 1-5 for additional primary antibodies. During various wash steps, handle the cells very carefully to minimize potential artifacts.

Option #2: Simultaneous incubation with unlabeled primary antibodies
This method is useful when the primary antibodies are from different hosts. For example, a mouse monoclonal antibody against antigen-X and rabbit polyclonal antibody against antigen Y.

1. After the blocking step, incubate the cells with unlabeled primary antibodies in the blocking buffer in a humidified chamber for one hour at room temperature or overnight at 4°C.


Tip: The blocking buffer can be prepared by mixing the serum from each host of the secondary antibodies. Alternatively, 5% BSA in PBS-T can be used as a blocking buffer.


2. Decant the antibody solution and wash the cells three times in PBS-T (5 minutes for each wash).


3. Incubate the cells with both secondary antibodies in 1% BSA for one hour at room temperature in the dark.


Tip: The secondary antibodies often come with a broad range of working dilutions. It is recommended to choose the dilutions very carefully and to employ additional optimization to see which dilution combination gives the best possible antibody staining.


4. Decant the secondary antibody solution and wash three times with PBS for five minutes each in the dark.


Tip: Additional washes may be required for a cleaner staining.

Option #3: Simultaneous incubation with directly labeled primary antibodies
This method is useful when the primary antibodies are from the same host. For example, a mouse monoclonal against antigen-X and a mouse monoclonal against antigen-Y.

1. After the blocking step, incubate the cells with directly labeled primary antibodies in the blocking buffer in a humidified chamber for one hour at room temperature or overnight at 4°C in the dark.


Tip: The use of fluorescently labeled phallodin is a good alternative to immunostaining actin as a reference marker in ICC.


2. Decant the antibody solution and wash the cells three times in PBS-T (five minutes for each wash).


Tip: Additional optimization may be required to determine the working number and time of washes.

COUNTERSTAINING, MOUNTING AND IMAGING

1. When all the necessary washing steps have been completed, cell nuclei can be counterstained with DAPI or Hoechst (1-10 µg/mL).


2. Invert the coverslip onto a glass slip with a drop of mounting media containing a fluorescence anti-fade agent.


Tip: If a coverslip of cells is accidently dropped and “cell side up” orientation is lost, you can still salvage the experiment by carefully picking up the coverslip with tweezers. Additionally, gently scraping one surface of the coverslip with a pipette tip is helpful to see if any cells are visibly removed.


3. Carefully remove the excess mounting media, if necessary, and seal as required with nail polish.


Tip: Some mounting media solutions have DAPI already added and will harden after exposure to air, eliminating the need to seal the edges of the coverslip.


4. Examine the cells under a fluorescence microscope and image as required.


Tip: Avoid long exposure to the excitation wavelength of the fluorochrome to prevent photo-bleaching. The slides may be stored in dark at -20°C or +4°C for re-examination.

POST-ANALYSIS DOUBLE IMMUNOFLUORESCENCE TIPS

1. Increase the percentage of blocking agents used in the blocking step and in the primary and secondary antibody dilution buffers. Increase the time of blocking and/or reduce the time of antibody incubations.


2. Dilute the amount of primary and/or secondary antibody.


3. Try quenching the fixation agent to eliminate any free aldehyde groups which could non-specifically bind antibody. Quench formaldehyde with 0.1M Tris or Glycine buffer and quench glutaraldehyde with 0.1% sodium borohydride.