Immunohistochemistry (IHC)

Immunohistochemistry (IHC) is a well-known staining procedure that uses antibodies to detect and localize proteins of interest, such as antigens, within the context of tissue structure. The technique is widely used as an ancillary method in clinical diagnostics to localize and quantify abnormal protein expressions linked to various diseases, particularly cancer. With specific tumor markers, researchers can determine tumor malignancy, stage, and grade, as well as identify and classify metastatic carcinomas of unknown primary sites. In basic research, IHC is used to study normal tissue and organ development, tissue repair, wound healing, and explore and validate critical biomarkers. While there are many variations of IHC staining, including IHC-frozen, IHC-paraffin, and IHC-free floating, the workflow for each method is similar, consisting of two parts: sample preparation and sample staining.

Sample Preparation

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Sample preparation is paramount to producing high-quality staining in IHC. When performed correctly, it will maintain cell morphology, tissue architecture, and the antigenicity of target epitopes during the experiment. Sample preparation involves several steps, including fixation, dehydration, embedding, sectioning, and antigen retrieval.

Tissue Fixation

Tissue fixation is responsible for preserving tissue morphology and the antigenicity of target molecules. Fixation is done using either formaldehyde- or alcohol-based fixatives, and the type of fixative used is influenced by the target antigen to be stained and the desired detection technique (i.e., fluorescence or chromogenic). Formaldehyde is the most widely used fixative for preserving protein targets. It can be used in perfusion and immersion fixation techniques for any length of time. However, the duration and other fixation conditions should be carefully optimized as they can impact preservation and tissue integrity. For instance, overfixation with formaldehyde can mask the epitope and produce strong non-specific staining, while underfixation may reduce or abolish tissue immunoreactivity. Following fixation in formaldehyde, tissue samples are typically embedded in paraffin wax before sectioning and further processing.

The most common alcohols used for tissue fixation are ethanol and methanol. When added to a sample, alcohol fixatives disrupt protein hydrogen bonds, altering their tertiary structure and water solubility, causing the proteins to precipitate. Compared to formaldehyde-based fixatives, alcohols are better at the preservation of antigenicity. They are more suitable for detecting membrane-bound proteins and studying DNA, RNA, and post-translational modifications such as phosphorylation. Because alcohols do not penetrate as well as formaldehyde, they are primarily used to fix frozen tissue samples after sectioning.

Tissue Embedding, Sectioning, and Mounting

Embedding is an essential part of IHC as it preserves tissue morphology and provides the tissue support during sectioning. Tissues that have been formaldehyde-fixed are usually dehydrated, embedded in paraffin wax, and sectioned into 4 to 5 µm slices using a microtome. These slices are then mounted onto glass slides coated with tissue adhesives (3-aminopropyltriethoxysilane (APTS), poly-L-lysine), dried overnight in an oven or microwave, and stored at room temperature for long periods. Before staining, formalin-fixed, paraffin-embedded (FFPE) tissues must be de-paraffinized, and an antigen retrieval step should be performed to recover antigenic sites that were masked during fixation.

Tissue samples that are too sensitive for chemical fixation or de-paraffinization can be snap-frozen by immersing the tissue in liquid nitrogen or isopentane. This is particularly useful for preserving proteins in their native state and detecting post-translational modifications. Tissues that have been frozen are sectioned into 5 to 20 µm thick slices on a cryostat and mounted on adhesive-coated slides (gelatin, poly-L-lysine). Frozen tissue slides can be safely stored at -80 °C for 6 to 12 months. On the day of the experiment, fix frozen tissue sections in alcohol for ten minutes before staining. Since alcohol fixatives do not mask epitopes, antigen retrieval steps are not required.

	Paraffin-embedded tissue	Frozen tissue
Fixation	The tissue sample is fixed in formaldehyde before being embedded in paraffin wax.	The tissue sample is snap-frozen by immersing in liquid nitrogen or isopentane and then fixed in alcohol.
Sectioning Instrument	Microtome	Cryostat
Storage	Stable for multiple years at room temperature	Store at -80 °C for 12 months
Advantages	Preserves tissue morphology Cost-effective, ambient storage conditions Best suited for tissue-archiving	Quicker protocol does not require de-paraffinization or antigen retrieval steps Preserves enzyme and antigen function Does not require an initial fixation step like FFPE tissues
Limitations	The procedure is laborious and time-consuming Proteins become denatured	Poor morphology Decreased resolution at high magnification Special storage requirements

De-paraffinization and Epitope Retrieval

The de-paraffinization process removes paraffin-embedded into the tissue and rehydrates the tissue with an aqueous solution in preparation for IHC staining. Failure to remove paraffin in FFPE sections will prevent antibodies from reacting with their target antigens, resulting in poor quality stains. To de-paraffinize FFPE tissue, slides are first heated to 55 °C for ten minutes to melt the wax. Next, several wash steps are performed to remove paraffin and rehydrate the tissue. These include:

• Multiple washes with xylene to dissolve and remove paraffin
• Graded washes with xylene and ethanol to remove xylene
• Rehydration of the sample through graded concentrations of ethanol in water
• A final rinse in pure water to remove ethanol

After de-paraffinization, antigen retrieval methods are used to recover antigenic epitopes masked during formaldehyde fixation. The two antigen retrieval methods for FFPE tissue include heat-induced epitope retrieval (HIER) and proteolysis-induced epitope retrieval (PIER). HIER is the most common method of antigen retrieval. It requires heating the slides to 95 °C for several minutes in a pre-heated retrieval solution, such as citrate (10 mM citric acid, 0.05% Tween 20, pH 6.0), EDTA (1 mM EDTA, 0.05% Tween 20, pH 8.0), or TBS (50 mM TBS, 0.05% Tween 20, pH 9.0) buffer. After heating, slides are cooled to room temperature and then gently rinsed with deionized water and PBS before staining. Various heating instruments can be used in HIER, such as a hotplate, microwave, pressure cooker, oven, or vegetable steamer. Each has its variation to the workflow mentioned. Therefore, optimal HIER conditions should be determined by the individual investigator.

The PIER method used proteases, such as trypsin (0.05%) or proteinase K (20 µg/mL), to partially digest proteins and unmask antigenic epitopes. This method requires careful optimization of the final enzyme concentration, the incubation temperature, and the incubation time. Failure to optimize these conditions can significantly lower the success rate for restoring immunoreactivity and destroy tissue morphology and antigen integrity.

Tissue Sample Labeling

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Tissue labeling is a multi-step process that uses antibodies to detect and visualize target antigens. It involves quenching the activity of endogenous proteins, blocking non-specific binding sites, immunodetection of the target protein, counterstaining (optional), and signal generation. Optimization at each step is critical in order to maximize signal detection and reduce background interference. Investigators must carefully consider their labeling strategy (direct or indirect), detection method (chromogenic or fluorescence), and the reagents needed to execute their experimental design.

Quenching Endogenous Protein Activity

Tissues contain several endogenous proteins such as peroxidases, phosphatases, and biotin. If not properly managed, the activity of these proteins may interfere with antigen detection and the staining process, resulting in false positives and significantly high background interference. Before incubating the tissue sample with the primary antibody, investigators must first determine which endogenous components will interfere with signal detection and then eliminate the interference using the appropriate blocking reagent. Determining which endogenous component to quench is ultimately linked to the type of tag conjugated to the signal generating antibody. For example, protocols that use the enzyme tag horseradish peroxidase (HRP) may require blocking with hydrogen peroxide to inhibit endogenous peroxidase activity and enhance sensitivity. The following table outlines methods commonly used to quench endogenous peroxidases, phosphatases, and biotin interferences.

If Using	Block	Tissue Types	Blocking Method
Biotinylated antibodies	Biotin	Liver, kidney, heart, brain, lung, mammary gland, and adipose tissue	To block endogenous biotin, first, incubate the tissue sample with streptavidin. Since streptavidin is tetrameric and can bind up to four biotin molecules, a subsequent incubation step with free biotin is required to block the remaining unoccupied biotin-binding sites.
HRP-labeled antibodies	Peroxidases	Liver, kidney, and vascular areas with red blood cells	Tissues can be tested for endogenous HRP by incubating with ABTS or TMB prior to primary antibody incubation. If a blue-green or blue color is observed, respectively, HRP is present, and blocking is required. Incubate tissue sections in 3% hydrogen peroxide for 10 minutes to quench endogenous peroxidase activity. Then wash three times with PBS. For delicate samples, where 3% hydrogen peroxide may damage the section or alter epitope accessibility, try a lower concentration of 0.3%.
AP-labeled antibodies	Phosphatases	Intestine, kidney, osteoblasts, lymphoid tissues, and placenta	Tissues can be tested for endogenous AP by incubating with substrate NBT/BCIP prior to primary antibody incubation; AP is present if a blue color is observed, and blocking is required. Quench endogenous phosphatase activity using 1 mM Levamisole; this is added with the chromogenic substrate. For intestinal AP, Levamisole has no effect. Block intestinal AP with 1% acetic acid before primary antibody incubation.

Blocking Non-Specific Interactions

The same attractive forces that govern antibody-antigen detection, such as hydrophobic interactions, ionic interactions, and other intermolecular forces, can also cause antibodies to weakly bind nonspecifically to sites that mimic the correct binding site on the target antigen. If not adequately managed, non-specific interactions can result in high background interference preventing visualization of the antigen of interest within its tissue architecture. Common causes of non-specific binding include interactions of antibodies or other detection reagents with serum proteins and ionic interactions between antibodies and tissues.

To mitigate non-specific interactions, a blocking step should be performed after sample preparation and just before incubation with the primary antibody. In general, any protein that does not bind to the antibodies or the target antigen can be used to block non-specific hydrophobic interactions. Common examples include bovine serum albumin (BSA), gelatin, non-fat dry milk, or serum from the same species as the secondary antibody. If staining with multiple secondary antibodies, blocking serum against all used secondaries is required. Blocking buffers containing BSA, gelatin, or non-fat dry milk added at 1-5% (w/v) final concentrations, are typically included in the diluents for the primary and secondary antibodies. In addition, non-ionic detergents including 0.3% Triton X-100™ or Tween 20™ can also reduce non-specific hydrophobic interactions.

Immunolabeling Tissue Samples

Detection of the target antigen with antibodies can be done using direct or indirect detection methods (figure 1). Determining which detection method to use depends upon the expression level of the target antigen, its accessibility, and the type of readout desired. Nevertheless, optimization at each step will help maximize signal detection.

Direct detection methods are commonly used to detect highly expressed antigens. The primary antibody against the target antigen is directly conjugated to a reporter enzyme (e.g., HRP or AP) or a fluorophore to facilitate detection and visualization. Enzyme-labeled antibodies are typically used for chromogenic detection and require subsequent incubation with an appropriate substrate for signal generation, such as TMB, ABTS, or ReadiUse™ StayRight™ Purple a safer non-mutagenic alternative to DAB. The reaction between the substrate and enzyme leads to the precipitation of insoluble, colored precipitates at the antigen localization site that can be visualized under a light microscope. For fluorescence detection, the primary antibody is conjugated to a fluorophore, such as iFluor® dyes and visualized under a fluorescence microscope.



In indirect detection, both primary and secondary antibodies are used to detect the target of interest. The primary antibody, which is unconjugated, binds directly to the target, while the secondary antibody, which is conjugated to a reporter enzyme or fluorophore, binds to the primary antibody. The labeled secondary antibody is typically directed against the immunoglobulin class or subclass of the primary antibody's species. For example, primary antibodies raised in rabbits will require an anti-rabbit secondary antibody raised in a host species other than a rabbit (e.g., goat anti-rabbit secondary). A major advantage of indirect detection is its ability to amplify the detection signal, which is made possible by the binding of multiple secondary antibodies to a single primary antibody. This significant increase in sensitivity makes indirect detection methods best suited for detecting low abundance targets. Other signal amplifying indirect detection methods compatible with IHC include PSA™ Signal Amplification, the Avidin-Biotin Complex (ABC), and the Labeled Streptavidin Biotin (LSAB) method.



Unlabeled primary antibodies and secondary antibodies, their respective conjugates, and enzyme substrates can be ordered directly from AAT Bioquest®. In addition, it is possible to directly label an antibody by using a commercially supplied labeling kit such as our ReadiLink™ Rapid Antibody Labeling Kits for fluorophore conjugations or our Buccutite™ HRP and Poly-HRP Antibody Labeling Kits.

Available Primary Antibodies	Available Secondary Antibodies by Target Species	Available Chromogenic Substrates
CD Antibodies	Anti-Rabbit Secondary Antibodies Anti-Mouse Secondary Antibodies Anti-Human Secondary Antibodies Anti-Donkey Secondary Antibodies	HRP ReadiUse™ ABTS Substrate HRP ReadiUse™ TMB Substrate HRP ReadiUse™ StayRight™ Purple Substrate StayRight™ Purple HRP Staining Kit pNPP Substrate for AP

Counter Staining

After labeling the target antigen using antibodies, a second chemical stain is often applied to the section to highlight specific tissue structures and add contrast to the primary stain. When selecting a counterstain, it is important to choose one that does not spectrally overlap with the primary stain. This will minimize the bleed-through effect and improve the discernibility of each signal. Single nuclear counterstains such as hematoxylin, nuclear fast red, and methyl green are commonly used for chromogenic detection methods. For fluorescent detection methods, target-specific fluorophores are used to counterstain nuclei, cytoskeletal components, amyloid deposits, and other tissue structures. These include DNA-binding dyes, such as DAPI, Hoechst, and Nuclear LCS1/DCS1™ dyes (see Table 1 below), phalloidin conjugates for staining F-actin, and amyloid stains like Congo Red.