Immunohistochemistry (IHC) Guide

Why choose IHC/ICC?

Immunohistochemistry (IHC) is an ideal technique to choose when experimental aims revolve around target co-localization with other proteins of interest, subcellular target localization, and expression profiles in different cell-cycle or cell-type subpopulations. It therefore has distinct advantages over related techniques, such as western blots and enzyme-linked immunosorbent assays (ELISAs).

Table 1

Key applications and differences between common biochemical techniques for protein detection.

History of IHC

One of the major early discoveries that facilitated the development of these techniques was the identification of the antibody-antigen interaction. Paul Erlich, a German scientist with a background in biochemistry and histopathology, won the Nobel Prize in physiology and medicine in 1908 for his work on this subject.³ One of the first immunostains, which consisted of anti-typhoid or anti-cholera antibodies conjugated to a red dye via tetrazotized benzidine, was created in the 1930s by John Marrack.⁴

Building on this work, Albert H. Coons developed a fluorescently conjugated antibody in the 1940s that closely resembled what we typically use today. His system consisted of a fluorescein-labeled anti-pneumococcal antibody, which could detect bacteria engulfed by macrophages.³ Another major advance took place in the 1960s with the development of enzyme conjugated antibodies, and further technical optimizations, such as the development of antigen retrieval and secondary antibody methods, occurred throughout the 1970s and 1980s.⁵

Coons, reflecting on his work, noted, “Fluorescent antibodies, whatever their scientific merits, are very attractive under the microscope. They shine in the dark, a brilliant greenish-yellow glow. Like pebbles in the moonlight, they weave a pattern in the forest which leads the weary children home”.³ Indeed, IHC and ICC are now essential methods in biological research and clinical diagnostics.

IHC workflow

The workflow of IHC and ICC are similar; the main steps are illustrated in Figure 2, alongside variables that must be taken into account at each stage to determine the ideal experimental conditions. Broadly, samples are isolated and chemically fixed to preserve cell and molecular organization, blocked to prevent non-specific interactions between the antibodies and the sample, and incubated with antibodies that recognize a target protein. Finally, the labeled proteins are visualized using chromogenic or fluorescent methods, and observed using microscopy.

Figure 2: Generalized protocol for IHC

After fixation, immunostaining incubation steps should be performed in a humidity chamber, particularly long incubations such as primary antibody incubation. A humidity chamber can be made using a plastic box with a lid and wet paper towels layered at the bottom of the box. To create raised platforms for the slides to sit on for ease of access during immunostaining, pipette tubes can be cut to the width of the box and taped in pairs to the bottom of the box.

Sample fixation and preparation

In order to visualize proteins in a tissue sample, that tissue must first be fixed and prepared to aid in sample handling and to make the tissue amenable to antibody penetration and detection.

Fixation is an essential step in IHC, because it preserves tissue integrity and morphology, prevents cellular degradation, and maintains the antigenicity of the target molecules. Sample preparation goes hand-in-hand with fixation, and will determine how the tissue will be sectioned (if at all) and handled, which fixatives are suitable, which kinds of visualization and imaging are possible, and how the tissue is stored.

Types of fixative

The choice of fixing solution will depend on sample type, antigen, and sample preparation; there is no ideal fixative for all tissues and situations. Though many fields have well-established standard fixatives that can serve as a starting point, it is worth experimenting with other fixatives when troubleshooting IHC. The optimal fixation conditions for a given experiment, length and temperature of fixation, and pH of the fixative, must ultimately be determined empirically to eliminate underfixation or overfixation. While underfixed tissues may undergo proteolytic degradation and destruction of the target epitope, overfixation and excessive cross-linking can mask the epitope or result in high non-specific background staining.

1. Formaldehyde

Formaldehyde-based fixatives create methylene (CH2) cross-links between proteins or between proteins and nucleic acids, and are commonly used due their strong tissue penetration and low background. Overfixation with formaldehyde can mask target epitopes by cross-linking the amino acids involved in their structure, although antigen retrieval techniques can resolve this problem.

2. Glutaraldehyde

Glutaraldehyde is a dialdehyde molecule and acts as a stronger crosslinker than formaldehyde. Crosslinking occurs quickly, but penetration of tissue by glutaraldehyde occurs slower than with formaldehyde due to the larger molecular structure. It is often used for electron microscopy, though rarely for IHC. In part this is due to its poor compatibility with paraffin embedding, and because it can produce high autofluorescence for IF. Tissues fixed with glutaraldehyde must also be treated with an aldehyde quencher such as ethanolamine before performing IHC, because otherwise free aldehyde groups can react with antibodies. Glutaraldehyde:formaldehyde mixtures can be used for fixation, benefitting from the rapid penetration of formaldehyde and the thorough fixation of glutaraldehyde.

3. Precipitative fixatives

Alcohol-based fixatives, such as methanol and ethanol, are structurally similar to water molecules, allowing them to participate in hydrogen bonding in fixed tissues. These fixatives cause proteins to precipitate by changing their dielectric points, and help stabilize proteins' secondary structure. However, alcohol fixatives do not preserve cell and tissue morphology as well as formaldehyde fixatives. Furthermore, antigen retrieval is usually not compatible with alcohol fixatives, meaning not all antibodies will work with alcohol fixation (Figure 3). Acetone can also be used to precipitate proteins in target tissues and cells, and is usually accompanied by fixation with formaldehyde or alcohols.

Figure 3: Effects of different fixatives on IHC outcome.

Staining of insulin (top) is mostly abolished following fixation of paraffin-embedded pancreas in ethanol (left) compared to formalin (right), while somatostatin staining is unaffected. Nuclei are counterstained with methyl green. Edited and reproduced under Creative Commons 4.0 CC-BY from Tsutsumi, Y. Pitfalls and Caveats in Applying Chromogenic Immunostaining to Histopathological Diagnosis. Cells 10, 1501 (2021).

4. Perfusion vs immersion

In perfusion fixation, fixative is injected through the vascular system of the still-intact organism, allowing for rapid fixation of organs and reducing non-specific staining of blood vessels. However, this method is technically challenging. Alternatively, the target tissue can be dissected and immediately immersed in fixative; this is usually appropriate for pieces of tissue less than 10 mm in size, and sufficient time must be allowed for full fixation, typically 2 – 24 hours at room temperature.

Sample preparation

The most appropriate sample preparation approach will depend on the downstream applications of an IHC experiment, as tissue processing affects how the sample will be handled, mounted and imaged, as well as the kinds of visualization and analysis that are possible. The specific details of sample preparation will vary greatly for different sample types and species, and must be determined empirically.

Paraffin embedding

Tissues that have been embedded in paraffin wax are stably preserved at room temperature for many years, making paraffin-embedding one of the most common methods for sample preparation of tissues. Paraffin-embedded tissues can be cut into thinner sections than frozen tissue (~5 μm versus ~10 μm, respectively), which can provide improved image quality and more accurate representations of tissue morphology during microscopic analysis.

Before tissue is embedded in paraffin, it is typically fixed in formalin for 4 - 24 hours to prevent cell death and changes in cell structure. The fixation step may be performed through perfusion or immersion. Following fixation, the tissue may be stored in 70% ethanol until paraffin embedding can be performed. Before embedding, the tissue sample must be dehydrated through immersion in increasing concentrations of alcohol, as paraffin wax cannot mix with water. After dehydration, the ethanol remaining in the sample is removed using xylene, and the sample is embedded in paraffin heated to ~60°C. Finally, tissue sections as thin as 4 μm are cut from the paraffin block using a microtome.

Frozen tissue

Frozen tissue samples are beneficial when detecting highly labile post-translational modifications or endogenous enzymatic activity. For example, alkaline phosphatase activity can be detected in frozen, acetone-fixed murine hair follicles7. Freezing tissue also has the advantage of significantly shorter processing times than paraffin-embedded samples.

Frozen tissue samples are typically prepared using liquid nitrogen or dry ice, and can be directly embedded in optimal cutting temperature (OCT) compound to facilitate cryosectioning. The frozen blocks of tissue may be used immediately, or stored at -80°C for up to 1 year. During cryosectioning, slices of tissue are directly placed onto microscope slides, and then immersion fixed. Depending on the intended target protein, a short (~20 minutes) alcohol or formaldehyde-based fixation may be performed. As a result, antigen retrieval is usually not necessary when using frozen tissue.

While frozen tissue is more advantageous than paraffin-embedded tissue in some ways, it cannot be used for long-term sample preservation, for which paraffin embedding is suitable. This is because ice crystals can form in the samples over long storage, leading to cell damage. In addition, the retention of endogenous enzymatic activity in frozen tissue sections can also present a problem for the detection step of immunostaining, meaning an additional blocking step may be required.

Fixed, free-floating sections

Tissues that have been embedded in paraffin wax are stably preserved at room temperature for many years, making paraffin-embedding one of the most common methods for sample preparation of tissues. Paraffin-embedded tissues can be cut into thinner sections than frozen tissue (~5 μm versus ~10 μm, respectively), which can provide improved image quality and more accurate representations of tissue morphology during microscopic analysis.

Fixed tissue can also be sectioned without paraffin embedding, particularly if thicker (>20 um) sections are taken and 3D structure of a tissue is important to the experimental aims. Fixation can either be by perfusion or immersion, which is followed by a cryopreservation step of incubating the tissue in 20-30% sucrose. Sucrose helps prevent ice crystal formation in tissues, protecting the tissue during freezing while sectioning. Sections are taken either on a cryostat or microtome, facilitated by embedding completely or partially in OCT, and are deposited into wells containing PBS. Once sections have been taken, the normal IHC protocol can be followed by sequentially transferring sections between wells containing the block, stain and wash solutions. Upon completion, the sections can be carefully moved onto slides using a paintbrush. Free-floating sections provide more 3D information, but are not suitable for long-term storage, and imaging multiple layers of cells through a thick section requires optical sectioning on a microscope to generate clear images.

Note that much thicker sections taken on a vibratome can also be used, which often involves embedding tissue in agarose before sectioning. For thick sections like this, much longer antibody incubation times may be necessary for adequate antibody penetration, and clearing might be required to mitigate light scatter through thick tissue when imaging.

Wholemount tissue

Wholemount tissue refers to tissue that has not been sectioned, leaving cells of interest completely intact within their native histological context. Wholemount samples include mouse and chick embryos, organoids, whole Drosophila, and even entire organs or nuclei from adult organisms. Wholemounts benefit from faster preparation times (no sectioning), but other steps, such as fixation and antibody incubation, may need to be longer to allow adequate penetration of the tissue, though the overall procedure is the same. Visualization can also be more complicated, depending on the tissue in question. Flat tissues, such as layers of muscle, may fit between slide and coverslip without distorting morphology, but others will be too thick or have a specific shape, such as embryos. These can be placed in a buffer within a glass-bottomed dish and optically sectioned on a confocal microscope, while very thick samples can be embedded in agarose and imaged on a lightsheet microscope. The specific details will be highly tissue and context-dependent, and should be determined on a case-by-case basis.

Primary antibody staining

Primary antibodies are generated by the immune system of a host organism (e.g. rabbit, guinea pig) that has been repeatedly immunized with an antigen. They recognize the protein of interest, rendering the target identifiable amongst the cellular milieu. A high specificity antibody that recognizes only the target of interest is essential for successful IHC, as non-specific binding can result in high background or false positive results. The following factors should be taken into account when selecting an antibody:

1. Reactivity

Primary antibodies will recognize antigens only from certain species, which will be the species that the immunizing antigen was originally from, as well as closely related species. For example, an antibody that recognizes a mouse protein target will often also recognize the same protein in rat, but possibly not in fish. The key determinant of this is how similar the epitope sequence is between species, allowing researchers to predict if an available antibody will work in a non-validated species.

2. Optimal conditions

As with the blocking step, the optimal primary antibody incubation conditions will vary depending on the specific antibody, the tissue sample, and the fixation conditions, among other factors. Most antibody datasheets will provide a recommended dilution range, usually within 1 - 25 μg/mL. A datasheet may give recommended concentrations to use relative to a master stock, such as 1:500, instead of absolute concentrations in μg/mL. This is particularly common for polyclonal antibodies. A range of 1:100 - 1:10000 is common to use for IHC. A higher concentration will give stronger signal, but more background, while a lower concentration will result in lower signal and background. The important factor is the SNR, which will also vary with different concentrations (Figure 6).

Which concentration is right for your application depends on a number of factors, including:

1. Tissue type: Some tissues have a greater propensity for higher non-specific binding, or stronger autofluorescence in certain spectral ranges.
2. Staining protocol: In general, tissue samples should be incubated with primary antibody overnight at 4°C. Overnight incubation provides the antibodies with time to bind to their target antigen, while incubating at a low temperature will decrease non-specific background interactions. Shorter incubation times at room temperature may also be appropriate in some cases, particularly for antibodies that produce very low background.
3. Washing protocol: A greater number of longer washes with a mild buffer (e.g. PBS only) will reduce non-specific binding but have little effect on strong antibody-epitope interactions.
4. Shaking: For IHC of free-floating sections or wholemount tissue, gentle to moderate shaking is recommended during all steps to reduce background and enhance antibody penetration. For sections already on slides, this is not necessary (due to the thinness of the sections) nor recommended (in case it dislodges the sections from the slides).
5. Microscopy capabilities: Confocal microscopes exclude out-of-focus light, including non-specific signal, which will increase SNR in the final image. Modern fluorescence detectors within microscopes also exhibit less detector noise, which is noise generated from the electronics in the detector itself. Powerful lasers enable visualization of dim signals that may not be visible with weaker LEDs, halogen lamps or lasers.

3. Host and multiplexing

The host is the species of animal an antibody was raised in. This is particularly important to know when using indirect detection methods (see Detection methods), because secondary antibodies are directed against the host species of primary antibodies, as illustrated in Figure 7. Where possible, primary antibodies should not be from the same host species as the tissue under investigation, such as using mouse primary antibodies on mouse tissue, because secondary antibodies (e.g. goat anti-mouse) will recognize not only the primary antibody but other endogenous proteins too.

When performing multiplexed IHC, if two primary antibodies are used from the same host species, the secondary antibodies will recognize both primary antibodies, making them impossible to distinguish during visualization. This is further complicated by secondary antibodies recognizing other secondary antibodies, which is a risk when using primary antibodies raised in goat (e.g. donkey anti-goat secondary antibodies will bind both primary and secondary antibodies raised in goat).