Tissue Preparation:

When starting immunocytochemistry with a new antibody or new tissue I consider the following items:

1.) Fixation

2.) Antibody Concentration

3.) Detergent

4.) Controls


Proper fixation is critical to good staining and obtaining nice preservation of the tissue. When working with a new antibody or tissue one needs to test out different lengths of fixation time to determine which time works best for the antibody being used. Some antibodies stain better with short fixation times while other antibodies require longer times. Both over and under fixing the specimen will result in no staining at all.

For animals whose tissues are fixed by perfusion, I test out various post-fixation times (i.e. no postfixation, 30 mins, 2 hrs, 4 hrs, 6 hr, or overnight). For cells grown in culture I generally test fixation times of 30, 60, and 90 mins initially.

One must be careful to not under fix the tissue as well, since you will have problems with nice ultrastructural preservation and your stain will leach out over a much shorter period of time.

One needs to determine the type of fixative to use. Generally, 4% paraformaldehyde is a good starting fixative for most antibodies. However, some antibodies work better in 2% paraformaldehyde, or in paraformaldehyde containing some glutaraldehyde.


When using a new antibody one needs to do a dilution series initially to determine what concentration of the antibody gives specific staining with the lowest background possible. If you use too high a concentration of antibody, you often get alot of background staining and then specific staining is not as easy to detect. If the antibody is used at a very low concentration, you may see no staining at all. A dilution series which goes from a high concentration to a very low concentration will allow you to choose a concentration that gives specific staining with low background.


Detergent can be used in the diluent for the antibodies to help them penetrate into the tissue, if the epitope for the antibody is intracellular. If the epitope for your antibody is extracellular one should not use detergent unless you want to determine both the intracellular and extracellular locations of the antigen in question. Triton X-100 is a good detergent to try. One should also test various concentrations of the detergent to see what provides the best staining. For example, I often use 0.1 % Triton X-100 when working with perfusion fixed tissue. However, I have used in a range of 0.05 % to 0.3 % Triton X-100 for different antibodies. For cells grown in culture or for cryostat sections being stained on a slide I usually use 0.01 % to 0.05 % Triton X -100 since the cells are more fragile and the cryostat sections are more likely to come of the slide the higher the concentration of detergent used. Always determine if detergent is required for the antibody in question and what concentration gives the best specific staining with low background. Too much detergent can sometimes increase the background.

When starting out I often start with 4% paraformaldehyde and 0.1% Triton-X 100 (for perfusion fixed tissue) or 0.025% Triton X-100 (for cells grown in culture) and adjust these variables from there. Of course, this does not apply if other fixatives or detergents have been used successfully before by other investigators for the particular antibody being used. It is best to get the most information about the antibody that you can from literature or from the supplier.


Controls are essential. Controls can include pre-immune serum from the animal in which the antibody was raised (if available) or normal serum from the same species that the antibody was raised in. These should be used at concentrations similar to what the antibody is being used at. In addition, a no primary antibody control should be done to test whether your secondary antibodies are causing non-specific staining. If available, the antibody should be preabsorbed with the antigen to help ensure the antibody staining is specific for the antigen. If you are collecting your images on a confocal microscope, you must collect images for the control tissue at the same range of settings as your are using for the labeled tissue. This helps you to determine if there is non-specific staining.