Methods for Antigen and Subcellular Structure Detection by Immunocytochemistry


To prepare cells for fixation with organic solvents, such as acetone or methanol, it is usually best to dry the cells prior to fixation. Typically at the time of harvest the cells are washed three times in phosphate-buffered saline (with Ca2+ and Mg2+) and allowed to air dry (bone dry). Fixation of cells by paraformaldehyde is done with washed cells without the drying step.


Antigens and cells respond differently to fixation and, as a result, fixation of cells often must be optimized for the cells, subcelluar structures, and antigens being investigated.

    II.A. Quality of fixatives:

    While some cells and antigens seem to be relatively insensitive to the quality of organic solvents used as fixatives, for others, the quality of the reagents is of considerable importance. Usually acetone and methanol with the least amount of assayed water is the best. Often this means at least spectrophotometer grade. Paraformaldehyde is best made up fresh from unopened ampules stored under nitrogen.

    II.B. Acetone:

    The classical method for fixation of cells for immunofluorescence used acetone as a fixative at room temperature. Fixation times varied between 3 and 20 minutes. Typically, cells were grown on coverslips, treated as required by the experimental protocol, and harvested by washing the cells three times in PBS (with Ca2+ and Mg2+) and air drying at room temperature to 37oC for up to 20 minutes. After drying, the coverslip cultures were fixed in acetone as described above. Once fixed, the cells could be stored in the refrigerator for several weeks before immunofluorescence staining.

    II.C. Acetone-Methanol:

    Acetone is not noted as the most effective fixative, but it is gentler on some antigens than methanol, which is a better fixative. In an attempt to gain the advantages of both reagents, acetone and methanol were mixed 1:1 (vol:vol) and used at -20oC. Typical fixation times are about 10 minutes. Post fixation, the excess fixative is removed by gently blowing across the fixed cells. This method is often used for cells that otherwise wash off of coverslips or culture dishes during immunocytochemical staining.

    II.D. Methanol:

    Methanol has been used as a fixative for immunofluorescence staining, but is usually less satisfactory than acetone alone or acetone:methanol, Except under special circumstances methanol fixation is not recommended by the IDTOIC for detection of antigens by immunofluorescence.

    II.E. Paraformaldehyde:

    Paraformaldehyde (1-4%) is an excellent fixative, but does not permeabilize cells. When internal antigens are the targets of immunocytochemistry, then paraformaldyde-fixed cells must be permeabilized postfixation. Common permeabilizing agents include triton-x 100 (0.3%), digitoxin (), n-octylicl, and BD FACSperm2.


    III.A. Direct immunofluorescence staining:

    Direct immunofluorescence staining uses antibodies with appropriate specificities to which fluorochromes have been directly conjugated. These reagents, while expensive, have two major advantages. First, they are easier and quicker to use than non-conjugated primary antibodies. Second, the use of conjugated antibodies avoids the problems of secondary antibody staining when two or more primary antibodies of the same species or isotype are used (e.g., mouse monoclonal antibodies). Cells, after appropriate fixation, are usually moistened with PBS (with Ca2+ and Mg2+) and the excess PBS is removed by careful aspiration. Then, a small amount of the appropriate dilution of conjugated antibody is added to the cells and the coverslip or culture vessel is placed in a humidified chamber in a 37oC CO2 incubator. For conservation of valuable primary antibody reagents, a drop of antibody may be placed on parafilm or wax paper and a coverslip culture placed cell-side down on the drop. Care must be taken in removing the coverslip for removal or excess antibody to not dislodge the cells (slightly pinching the parafilm or wax paper and the use of fine-tipped Dumont forceps is recommended). Incubation times for the immunofluorescence labeling vary between 30 minutes and overnight. Following incubation, excess antibody is washed off with PBS and the cells are mounted in glycerol:PBS (1:1, vol:vol) or SloFade (Molecular probes). The OIC has found Slofade to be effective in enhancing the duration of the fluorescence signal.

    III.B. Indirect immunofluorescence staining:

    Indirect immunofluorescence staining is the most commonly used fluorescence staining procedure, principally because of the limited availability of conjugated primary antibodies. In this procedure the cells are first incubated with a primary antibody directed to a target protein. The procedure is initially identical to that described above for the direct conjugated antibody. After the reaction with the primary antibody is complete, then the excess antibody is removed by three sequential washes in PBS. Then, secondary antibody is applied and incubated in the same manner as the primary, except that incubation rarely exceeds 30 minutes at 37oC. Excess secondary antibody is removed by three sequential washes in PBS and the cells are mounted in glycerol PBS or SloFade (Molecular probes). Dilution series of the both the primary and secondary antibodies are usually necessary to optimize the immunofluorescence staining. In addition, conjugate controls and nonspecific primary antibody controls are necessary to monitor nonspecific fluorescence reactions.

    III.C. Non-antibody-mediated fluorochome labeling.

    A large number fluorochrome labeling procedures have been developed which do not depend on antibodies. These techniques are well described by the manufacturers and the OIC recommends that their instructions be followed carefully. For enzyme activity studies that are fluorescence-based (e.g., with coumarin-based substrates), we recommend that researchers discuss their requirements with the OIC before starting their work.

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