Immunohistochemistry

The tissue may then be sliced or used whole, dependent upon the purpose of the experiment or the tissue itself. Before sectioning, the tissue sample may be embedded in a medium, like paraffin wax or cryomedia. Sections can be sliced on a variety of instruments, most commonly a microtome, cryostat, or vibratome. Specimens are typically sliced at a range of 3 µm-5 μm. The slices are then mounted on slides, dehydrated using alcohol washes of increasing concentrations (e.g., 50%, 75%, 90%, 95%, 100%), and cleared using a detergent like xylene before being imaged under a microscope.

Depending on the method of fixation and tissue preservation, the sample may require additional steps to make the epitopes available for antibody binding, including deparaffinization and antigen retrieval. For formalin-fixed paraffin-embedded tissues, antigen-retrieval is often necessary, and involves pre-treating the sections with heat or protease.[1][5] These steps may make the difference between the target antigens staining or not staining.

Depending on the tissue type and the method of antigen detection, endogenous biotin or enzymes may need to be blocked or quenched, respectively, prior to antibody staining. Although antibodies show preferential avidity for specific epitopes, they may partially or weakly bind to sites on nonspecific proteins (also called reactive sites) that are similar to the cognate binding sites on the target antigen. A great amount of non-specific binding causes high background staining which will mask the detection of the target antigen. To reduce background staining in IHC, ICC and other immunostaining methods, samples are incubated with a buffer that blocks the reactive sites to which the primary or secondary antibodies may otherwise bind. Common blocking buffers include normal serum, non-fat dry milk, BSA, or gelatin. Commercial blocking buffers with proprietary formulations are available for greater efficiency. Methods to eliminate background staining include dilution of the primary or secondary antibodies, changing the time or temperature of incubation, and using a different detection system or different primary antibody. Quality control should as a minimum include a tissue known to express the antigen as a positive control and negative controls of tissue known not to express the antigen, as well as the test tissue probed in the same way with omission of the primary antibody (or better, absorption of the primary antibody).[1]

The antibodies used for specific detection can be polyclonal or monoclonal. Polyclonal antibodies are made by injecting animals with the protein of interest, or a peptide fragment and, after a secondary immune response is stimulated, isolating antibodies from whole serum. Thus, polyclonal antibodies are a heterogeneous mix of antibodies that recognize several epitopes. Monoclonal antibodies are made by injecting the animal and then taking a specific sample of immune tissue, isolating a parent cell, and using the resulting immortalized line to create antibodies. This causes the antibodies to show specificity for a single epitope.[6]

For immunohistochemical detection strategies, antibodies are classified as primary or secondary reagents. Primary antibodies are raised against an antigen of interest and are typically unconjugated (unlabeled), while secondary antibodies are raised against immunoglobulins of the primary antibody species. The secondary antibody is usually conjugated to a linker molecule, such as biotin, that then recruits reporter molecules, or the secondary antibody itself is directly bound to the reporter molecule.[3]

Reporter molecules vary based on the nature of the detection method, the most popular being chromogenic and fluorescence detection mediated by an enzyme or a fluorophore, respectively. With chromogenic reporters, an enzyme label reacts with a substrate to yield an intensely colored product that can be analyzed with an ordinary light microscope. While the list of enzyme substrates is extensive, alkaline phosphatase (AP) and horseradish peroxidase (HRP) are the two enzymes used most extensively as labels for protein detection. An array of chromogenic, fluorogenic and chemiluminescent substrates is available for use with either enzyme, including DAB or BCIP/NBT, which produce a brown or purple staining, respectively, wherever the enzymes are bound. Reaction with DAB can be enhanced using nickel,[7] producing a deep purple/black staining.

Fluorescent reporters are small, organic molecules used for IHC detection and traditionally include FITC, TRITC and AMCA, while commercial derivatives, including the Alexa Fluors and Dylight Fluors, show similar enhanced performance but vary in price. For chromogenic and fluorescent detection methods, densitometric analysis of the signal can provide semi- and fully quantitative data, respectively, to correlate the level of reporter signal to the level of protein expression or localization.

Chromogenic immunohistochemistry: The cell is exposed to a primary antibody (red) that binds to a specific antigen (purple square). The primary antibody binds a secondary (green) antibody that is chemically coupled to an enzyme (blue). The enzyme changes the color of the substrate to a more pigmented one (brown star).

The direct method is a one-step staining method and involves a labeled antibody (e.g. FITC-conjugated antiserum) reacting directly with the antigen in tissue sections. While this technique utilizes only one antibody and therefore is simple and rapid, the sensitivity is lower due to little signal amplification, in contrast to indirect approaches.[3] However, this strategy is used less frequently than its multi-phase counterpart.

The indirect method involves an unlabeled primary antibody (first layer) that binds to the target antigen in the tissue and a labeled secondary antibody (second layer) that reacts with the primary antibody. As mentioned above, the secondary antibody must be raised against the IgG of the animal species in which the primary antibody has been raised. This method is more sensitive than direct detection strategies because of signal amplification due to the binding of several secondary antibodies to each primary antibody if the secondary antibody is conjugated to the fluorescent or enzyme reporter.[3]

Further amplification can be achieved if the secondary antibody is conjugated to several biotin molecules, which can recruit complexes of avidin-, streptavidin- or NeutrAvidin protein-bound enzyme.[3] The difference between these three biotin-binding proteins is their individual binding affinity to endogenous tissue targets leading to nonspecific binding and high background; the ranking of these proteins based on their nonspecific binding affinities, from highest to lowest, is: 1) avidin, 2) streptavidin and 3) NeutrAvidin protein.

The indirect method, aside from its greater sensitivity, also has the advantage that only a relatively small number of standard conjugated (labeled) secondary antibodies needs to be generated. For example, a labeled secondary antibody raised against rabbit IgG, which can be purchased "off the shelf", is useful with any primary antibody raised in rabbit. This is possible because all rabbit IgG in this example would have the same Fc (constant) region, thus even with a small number of anti-rabbit secondary antibodies generated, each could adhere to any primary rabbit antibody.[3] This is particularly useful when a researcher is labeling more than one primary antibody, whether due to polyclonal selection producing an array of primary antibodies for a singular antigen or when there is interest in multiple antigens. With the direct method, it would be necessary to label each primary antibody for every antigen of interest.

After immunohistochemical staining of the target antigen, a second stain is often applied to provide contrast that helps the primary stain stand out. Many of these stains show specificity for specific classes of biomolecules, while others will stain the whole cell.[3] Both chromogenic and fluorescent dyes are available for IHC to provide a vast array of reagents to fit every experimental design, and include: hematoxylin, Hoechst stain and DAPI are commonly used.

Tumor biology allows for a number of potential intracellular targets. Many tumors are hormone dependent. The presence of hormone receptors can be used to determine if a tumor is potentially responsive to antihormonal therapy. One of the first therapies was the antiestrogen, tamoxifen, used to treat breast cancer. Such hormone receptors can be detected by immunohistochemistry.[13] Imatinib, an intracellular tyrosine kinase inhibitor, was developed to treat chronic myelogenous leukemia, a disease characterized by the formation of a specific abnormal tyrosine kinase. Imitanib has proven effective in tumors that express other tyrosine kinases, most notably KIT. Most gastrointestinal stromal tumors express KIT, which can be detected by immunohistochemistry.[14]

Many proteins shown to be highly upregulated in pathological states by immunohistochemistry are potential targets for therapies utilising monoclonal antibodies. Monoclonal antibodies, due to their size, are utilized against cell surface targets. Among the overexpressed targets are members of the EGFR family, transmembrane proteins with an extracellular receptor domain regulating an intracellular tyrosine kinase.[15] Of these, HER2/neu (also known as Erb-B2) was the first to be developed. The molecule is highly expressed in a variety of cancer cell types, most notably breast cancer. As such, antibodies against HER2/neu have been FDA approved for clinical treatment of cancer under the drug name Herceptin. There are commercially available immunohistochemical tests, Dako HercepTest, Leica Biosystems Oracle[16] and Ventana Pathway.[17]

Similarly, EGFR (HER-1) is overexpressed in a variety of cancers including head and neck and colon. Immunohistochemistry is used to determine patients who may benefit from therapeutic antibodies such as Erbitux (cetuximab).[18] Commercial systems to detect EGFR by immunohistochemistry include the Dako pharmDx.