IHC builds on histochemistry by specific localization of target components within cells and tissue. This is only possible if the cells and tissues are appropriately processed and fixed such that they are recognizable under the microscope. Different fixation and tissue processing methods may affect antibody recognition such that strong staining is possible with one method, but not with another. In general, for research purposes, antibody staining is easier in frozen sections; however, tissue architecture is better in paraffin sections.

Because devitalized tissues undergo autolysis from release of lysosomal proteolytic enzymes, preservation of antigenicity requires either rapid fixation, which inactivates enzymes, or placing the tissue at sufficiently cold temperatures where enzymes are inactive. Fixation serves two additional purposes, to immobilize molecules within the cell and prevent diffusion and to increase tissue rigidity to improve sectioning. Antigen localization is optimized by more rapid fixation, either by improved delivery with perfusion fixation or by acceleration of the chemical process using heat or agitation [20]. Perfusion fixation has the additional advantage of improved vessel morphology. Small animals can be easily perfused at physiologic pressures through either the heart or the aorta using a sphygmomanometer or gravity-based perfusion fixation apparatus (Fig. 10.3).

As previously mentioned, freezing specimens generally leads to loss of tissue architecture due to the formation of ice crystals. Two strategies can be used for preservation of antigenicity while minimizing loss of architecture. One is to freeze the sample as rapidly as possible, by immersing it in isopentane cooled to its freezing point using liquid nitrogen, placing it on a metal block cooled by liquid nitrogen, or immersing it in some combination of dry ice and either acetone or ethanol. The other strategy is to use cryoprotection and dehydration. Sucrose is commonly used as a cryoprotectant [20]. For research purposes, specimens are generally embedded and frozen in optimal cutting temperature compound (OCT, mixture of polyvinyl alcohol and polyethylene glycol) to allow frozen sectioning. Cell membrane antigens and cytokines are frequently more successfully stained with frozen section rather than with paraffin-fixed tissue.

Chemical fixatives can be roughly divided between those that cause protein cross-linking and those that cause protein coagulation or precipitation. The most commonly used cross-linking fixative is formalin solution (37–40 % formaldehyde existing as low polymers and 60–63 % water). The low polymers of formaldehyde must be broken down to monomeric formaldehyde for active fixation by protein cross-linking through the aldehyde group, also known as a methylene bridge. This can be done by dilution with a buffer solution at physiological pH, generally to 4 or 10 %. Methanol (10 %) is frequently added to formalin by the manufacturer to slow down polymerization to paraformaldehyde. Paraformaldehyde, which is commonly used for research purposes, consists of higher polymers of formaldehyde and requires heating in buffered solution to 60 °C within a fume hood for solubilization [21]. Currently, both methanol-free formalin and aliquots of frozen formaldehyde prepared from paraformaldehyde are also commercially available. Although formaldehyde generally penetrates the tissue after 12–24 h, significant cross-linking may require up to 7 days to occur. As increased cross-linking tends to decrease antigenicity, over-fixation may render specimens unusable for immunohistochemical analysis.

Glutaraldehyde has two active aldehyde groups that allow more thorough cross-linking, making it ideal for electron microscopy, but less ideal for IHC as the free aldehyde groups must be removed or blocked to prevent nonspecific binding, and the thorough cross-linking frequently destroys antigenicity. The more thorough cross-linking also impedes paraffin penetration. From a practical standpoint, the more expensive “electron microscopy (EM) grade” glutaraldehyde that contains the monomer and low polymer forms required for adequate cell and tissue penetration rather than the “technical grade” glutaraldehyde that has larger polymers should be used for research purposes [21].

Coagulating fixatives include acetone, ethanol, methanol, trichloroacetic acid, and zinc-based fixatives. Acetone, ethanol, and methanol when used alone can cause significant shrinkage and hardening artifacts but do allow better preservation of DNA and RNA as well as preservation of antigenicity without the need for antigen retrieval. Carnoy’s fixative (60 % ethanol, 30 % chloroform, 10 % glacial acetic acid) or methyl Carnoy is a commonly used non-formalin fixative and may preserve carbohydrate and cytosolic membrane antigens more successfully than aldehyde-based fixatives [20].

Whole mounts can be prepared for IHC without sectioning. They provide an immediate and three-dimensional view of stained antigens, which is useful for rapid surveillance. In general, the technique is limited to tissues that can be prepared as small blocks (<5 mm thick) such as embryos. Antibody penetration can be limited, however, resulting in patchy or false-negative staining.

Frozen tissue embedded in OCT may be cut immediately using a cryostat. Different tissues required cutting at different temperatures, which generally range between −20 and −30 °C. Slice thickness varies depending on the tissue and the skill of the operator but generally ranges between 5 and 10 μm. Thinner sections provide less possibility of cellular overlap and improved ultrastructural resolution. Certain applications, such as confocal microscopy, may call for considerably thicker sections. Cutting may introduce folds or wrinkles in the tissue, degrading its stainability and architecture. A newer method of tape transfer for frozen sections has been developed to improve tissue architecture. Essentially, tape is applied to the block prior to sectioning. After the section is cut, it is transferred tissue side down to a polymer-coated slide. The tissue is bonded to the slide using a flash of ultraviolet light, at which point the tape is removed, and the slides can then be processed as regular frozen sections for staining [22]. Tape transfer systems are commercially available for purchase, but the added expense may or may not be justifiable depending on the goals of the experiment. Slides should be maintained at −20 or −80 °C after cutting to maintain antigenicity.