The Typical Immunohistochemistry Workflow

blog / Pathology February 03 2020

Immunohistochemistry (IHC) ties together immunological, biochemical and microscopic techniques to yield visual information about proteins and other macromolecules in tissues. Targets are detected through the use of labelled antibodies that penetrate tissues and specifically bind proteins and other cellular components of interest, e.g., organelles.

Unlike other immunoassay formats such as western blotting and ELISA, which provide qualitative and quantitative information about target molecules that are removed from their normal biological location, IHC allows us to qualitatively and quantitatively analyse target molecules in situ, thus providing information about these components in their normal (or diseased) histological context. There are many approaches and variations within IHC, depending on the setup and experimental goal. In this article, we will take you through a typical IHC workflow.

Figure 1: Typical Immunohistochemistry Workflow

1. Tissue Preparation

Proper tissue handling and preparation is critical for a reliable staining outcome. Here, epitopes and tissue morphology are typically preserved in formalin-fixed paraffin-embedded (FFPE) tissue blocks.

Tissues (e.g. biopsies) or organs of interest are first fixed in formalin*, which chemically cross links the proteins in the tissue, locking all cellular processes, proteins and macromolecules conformationally in their exact location at the time. After fixation, the tissues are embedded in blocks of paraffin wax and then sectioned into very thin slices approximately 4-5 µm thick using a microtome. Sectioned tissues are mounted onto glass slides that are coated with a tissue adhesive. The slides are then dried prior to further processing. For best results, it is wise to use freshly isolated biopsies or small pieces of tissue, and fix them immediately after collection to preserve tissue architecture and prevent degradation. A small number of proteins will become degraded during the FFPE sample workflow. One solution to staining such proteins by IHC is to cryopreserve the sample immediately after tissue dissection or after a mild fixation, typically in an alcohol solution. Frozen samples are then embedded in blocks of a water-soluble resin, and sectioning is carried out in a cryostat.

*Formalin is not the only fixative used in IHC. Depending on the sample type and sensitivity of target epitopes, other aldehydes, alcohol solutions or freezing in a cryoprotective solution may also be used.

2. Epitope Retrieval

Formalin-induced cross links or long-term fixation in general may mask certain epitopes, preventing antibody binding in subsequent steps. The goal of epitope retrieval is to unmask or re-expose these epitopes. This is usually carried out after a pretreatment step to rehydrate the sample and completely remove all paraffin so that it doesn’t interfere with staining in downstream steps. A number of strategies may be used here. Treatment with digestive enzymes, heat or detergents or a combination of these three is common. The most popular approach to epitope retrieval in FFPE samples is heat induced epitope retrieval (HIER) in a Tris-EDTA buffer at pH 9. Bear in mind that while paraffin removal is essential prior to staining, epitope retrieval is not necessary for all epitopes. If there are no documented epitope retrieval guidelines for your target epitope(s), either in the literature or in antibody supplier datasheets, you will need to optimise this step yourself through trial and error.

3. Blocking Off-Target Binding

Non-target binding sites on proteins present in the tissue sample are blocked prior to primary antibody addition. This is achieved by incubation in a buffer containing serum, non-fat milk powder, bovine serum albumin or another blocking agent. This buffer sometimes includes a small amount of one or more gentle detergents, e.g., Tween-20, SDS, to aid in wetting.

It may also be necessary to block endogenous proteins that mimic or interfere with the proteins used in downstream detection systems (e.g., biotin, avidin, horseradish peroxidase) to reduce background and the risk of false positive detection. This is generally done by physically or chemically blocking all endogenous biotin or enzyme activity.

4. Primary Antibody

Here, a primary antibody specific for the target epitope is applied to the slide*. Primary antibodies for IHC should be highly specific and penetrate the target tissue. Less specific antibodies are likely to bind off-target epitopes leading to background signals that may significantly impact your ability to interpret experimental results.

Monoclonal or polyclonal antibodies may be used here, each with their own advantages and drawbacks. In summary, polyclonal antibodies can bind many epitopes on the same target protein and are thus potent. However, the ability to bind multiple epitopes can lead to reduced specificity and off-target binding. Monoclonal antibodies on the other hand only bind a single epitope and are often more specific, but they may not be ideal for low-abundant targets. You can read more about this in our previous article about primary antibody selection for IHC.

No matter which type of antibody is used, it is necessary to optimise antibody concentration for every new assay (i.e. each time a new sample/tissue type is used). Careful optimisation of this step is critical in IHC because the success of a primary antibody is not only dependent upon its specificity and affinity for the target, but also on the abundance and availability of on and off-target epitopes in the sample. Using too much antibody may increase background if the target epitopes become saturated with bound antibody. Using less primary antibody may alleviate this issue, but with the possible tradeoff of a lower intensity signal, and in the case of a low-affinity antibody a false negative result may occur. Unbound primary antibody is removed by rinsing the slides in a suitable wash solution that usually contains trace amounts of detergent and salts.

*Other binding molecules besides antibodies may be used such as antibody fragments, peptides or other small molecules.

5. Detection

During detection, the primary antibody-target interaction is localised and visualised as a measurable signal. Although other methods exist, antibody-based detection methods are most widely used. These can be grouped into two categories: direct and indirect, depending on whether the primary (direct) or secondary antibody (indirect) is conjugated to an enzyme or fluorescent label. Selecting the right detection method requires considerations about the expected level of target expression, its accessibility and the type of readout required and equipment available, i.e. chromogenic signal (light microscopy) or fluorescent signal (fluorescence microscopy).

In direct detection, the primary antibody is labeled with an enzyme or fluorophore, and the signal generated upon substrate addition or excitation of the fluorophore becomes the assay readout.

Indirect detection methods often exploit the interaction between biotin and its binding partners e.g., strept(avidin) to increase detection sensitivity. Here, after washing away unbound primary antibody, a biotinylated secondary antibody is applied, and its presence is localised at the target site following the application of an avidin-enzyme complex (e.g., horseradish peroxidase or alkaline phosphatase), where the enzyme produces an insoluble coloured precipitate in the presence of a suitable substrate. Because each biotin molecule can bind multiple molecules of avidin, the detection signal is amplified. If fluorescence detection is required, a fluorescently-labelled avidin or enzyme is used to generate a fluorescent signal.

As mentioned earlier, endogenous biotin may lead to background issues in IHC staining. To overcome these issues, you can opt for a polymer-based detection method instead of a biotin-avidin setup. Here, the secondary antibody is directly linked to a polymer of enzymes instead of a single enzyme. Because each secondary antibody is linked to many enzymes, the signal from each primary antibody-antigen interaction is greatly amplified, thus increasing assay sensitivity.

6. Counterstaining

Once staining is complete i.e. after detection, it is often beneficial to include a counterstain to enhance the contrast between stained and unstained areas in the tissue section, making it easier to pinpoint interesting histological features. A number of counterstains exist, and some are cell structure specific. Haematoxylin is the most commonly used for FFPE samples when light microscopy-based detection methods are used. This stains cytoplasm pale blue and nuclei dark blue. When fluorescence detection is used, fluorescent dyes that selectively bind nucleic acids are usually used as counterstains to label cell nuclei.

After counterstaining, coverslips are applied to the stained slides which are then sealed, usually with clear nail varnish or an appropriate commercially available resin. This step is important to prevent solubilisation of the enzymatic products generated during detection, and to prevent photobleaching by any remaining fluorophores. The sealed slides are then suitable for image analysis and long-term storage.

7. Image Analysis

Depending on which detection format is used, the slides are visualised by light or fluorescence microscopy. One may also choose to perform confocal microscopy for greater detail and enhanced imaging capabilities. Additionally, samples can be analysed by high content screening for rapid quantitation and comparison of data from multiple samples. And with the right equipment, it is also possible to scan the slides to get high-quality images in digital format for further analysis and for use in publications and presentations.

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