A Guide to Better IHC Staining: Steps, Troubleshooting, and Optimization

Immunohistochemistry (IHC) is a strong and popular method in tissue studies and medical testing. It mixes body structure, immune system, and chemical techniques to find and show specific proteins in tissue pieces. This happens by using the tight connection between an antibody and its target protein. A marker, like an enzyme or glowing dye, makes the protein visible. Albert H. Coons and his team started IHC in the 1940s with glowing dyes. Since then, it has grown a lot with better ways to prepare tissues, unmask proteins, link antibodies, stain samples, and use microscopes.

IHC is a helpful extra tool. It works alongside older staining methods like Hematoxylin and Eosin (H&E) and Special Stains. H&E and Special Stains mostly show tissue shape and aren’t specific. But IHC targets certain protein markers. It gives key details about where they are, how they’re spread, and how strong they are in cells and tissues. This makes IHC a common tool in labs. It’s also super important in hospitals for sorting cells and diagnosing diseases. IHC helps identify illnesses, sort tumors, find where cancers spread from, spot tiny tumor spots, and give clues about treatment outcomes based on protein signs.

The IHC process has several main steps. These are split into preparing the sample and staining it. Knowing each step and making them work well is super important for clear, trustworthy results.

Sample Preparation

Getting the tissue ready is the first big step in IHC. It really affects how good the final staining looks. This step includes a bunch of actions to keep the tissue’s shape and the protein’s ability to be detected.

It starts with getting the tissue sample. This can be small pieces or whole organs from people or animals. Quick saving is key to stop proteins from breaking down and to keep tissue structure. Sometimes, the tissue is cleaned in vivo or in vitro to remove blood parts that might mess with protein detection.

Tissue Fixation

After collecting, the tissue gets fixed. This step links proteins together and keeps cell structure. The type of fixative, its strength, pH, and how long it’s used matter a lot. These can help or hurt staining quality. No fixative works perfectly for all proteins. Formaldehyde, often as formalin, is a common fixative that links proteins and can be partly undone. It’s used for soaking or flushing tissues. It’s popular for formalin-fixed paraffin-embedded (FFPE) tissues. But long or wrong fixation can hide target proteins. Other fixatives like acetone or methanol might be better for some proteins. For proteins that don’t like aldehyde fixation, quick freezing in liquid nitrogen and cutting with a cryostat, then fixing with cold acetone or alcohol, may be needed.

Fixation must be steady and known to give good results. Uneven fixation can cause spotty staining. Too little fixation (under 12 hours) or too much (over 48 hours) can lead to wrong results or faint staining. Dead tissue areas should be skipped. They might stain randomly or not at all.

Tissue Embedding

After fixing, tissues are placed in a material to hold their shape, allow long storage, and make thin slicing easy. Paraffin wax is the usual choice for regular tissue work, especially for FFPE samples. FFPE tissues give good results for many proteins, especially with protein unmasking methods.

But some proteins don’t handle the chemicals or heat of paraffin embedding well. In those cases, tissues can be frozen in a cold material and snapped frozen in liquid nitrogen. These frozen samples are cut with a cryostat. Vibratome slices are another option. They avoid harsh chemicals or high heat, keeping proteins active. But they’re slow to cut and might have rough marks.

Sectioning and Mounting

Once set in a material, the tissue block is sliced into thin pieces. FFPE tissues are cut at 4–7 μm thick using a microtome. These slices are placed on glass slides. Charged or sticky slides are best. They help tissues stick during staining. Sticky coatings like APTS or poly-L-lysine add gripping groups to slides. Poorly stuck or uneven slices can cause patchy staining and uneven background. Protein-based glues in the water bath for charged slides should be avoided. They can block the slide, causing weak sticking and pooling of staining liquids. Mounted slices are usually dried in an oven or microwaved.

Frozen slices are cut with a cryostat at cold temperatures. They’re also placed on sticky slides. These slices are often dried overnight at room temperature. Then, they’re fixed again, usually in cold acetone, fresh paraformaldehyde, or formalin. Frozen slices are great for sensitive proteins but might have worse shape and clarity than paraffin slices.

Deparaffinization and Epitope (Antigen) Retrieval

For FFPE slices, all paraffin wax must be removed before staining. This lets antibodies reach target proteins. Organic solvents like xylene are used. If wax stays, it hides proteins and pushes away water-based staining liquids.

Fixation, especially with formaldehyde, makes links (methylene bridges) that can hide protein parts. This makes antibody binding hard or impossible. Antigen or epitope retrieval (AR) is a key step for FFPE tissues. It breaks these links to show hidden protein spots. Finding and improving AR methods has made IHC much more sensitive and useful for FFPE samples.

There are two main ways for AR:

Heat-Induced Retrieval (HIER): This is the top method. It heats or boils slices in different buffer solutions with various pH levels. Common tools include microwaves, pressure cookers, autoclaves, or water baths. Citrate buffer (pH 6.0) is a favorite solution. Tris-EDTA (pH 9.0) and EDTA (pH 8.0) are also used a lot. Getting the right temperature, time, and buffer pH is super important for good results.

Proteolytic-Induced Retrieval (PIER): This uses enzymes like pepsin or trypsin to break down tissue and show protein parts. It’s used for proteins that might lose activity with heat. But it can harm tissue shape or even destroy the protein. So, HIER is usually better unless the antibody maker says otherwise.

Bad or missing AR can cause poor staining. A control slide without AR should be used to spot errors from the process. Adjusting AR for the antibody, tissue type, and fixation method is key.

Quenching/Blocking Background Activity

Many detection systems use natural enzymes or biotin in tissues. These can cause wrong positive signals or extra background. Steps must stop or hide these natural parts.

Common things to block include:

Natural Peroxidases: Most tissues have these. They can cause errors with HRP-linked antibodies. Blocking is a must for clear staining. This is often done with hydrogen peroxide solutions.

Natural Alkaline Phosphatase: These can mess with antibodies linked to this enzyme. Blocking with levamisole helps if this enzyme is used.

Natural Biotin: Found in tissues like liver, it can interfere with biotin-based detection. Blocking with biotin-saturated avidin systems can help.

Blocking Non-Specific Binding

Antibodies can stick to wrong spots in tissues. This can happen due to things like Fc receptors or sticky attractions. This causes extra background that hides the true signal.

Blocking non-specific binding is a big step before adding the main antibody. This is done with a blocking buffer. Common options include normal serum from the same animal as the secondary antibody or special blocking products. Examples are soy milk, egg whites, or store-bought mixes. Longer blocking time can reduce background. Avoiding extra staining from poor blocking is super important.

Sample Staining

After preparing and blocking the sample, staining starts. This involves adding antibodies and detection stuff to show the target protein. This multi-step process needs careful work at every part to get strong signals and low background.

Antibodies, both main and secondary, are mixed into a buffer. This keeps them stable, helps them spread, and stops unwanted sticking. Picking the right antibody and finding the best mix is key for clear results.

Antibodies (Primary and Secondary)

The main antibody goes first. It sticks to the target protein exactly. How well it works and how picky it is are super important. Testing it with a Western blot on known and unknown samples can show if it’s good for IHC.

Immunodetection (Detection Systems)

Showing the antibody-protein link needs a detection system. The choice depends on how much target protein is there and how strong the signal needs to be.

Different ways exist to mark antibodies or layers for seeing:

Direct Method: The main antibody is marked and put on the tissue. This is fast but not very strong because there’s no signal boost. It’s not used much.

Indirect Method: A marked secondary antibody sticks to the unmarked main antibody. This is stronger because it boosts the signal. One marked secondary antibody can work with many main antibodies from the same animal.

Detection systems can use glowing molecules or enzymes.

Glowing Methods: Use antibodies with glowing dyes (e.g., FITC, rhodamine, Texas red). These need a glowing microscope. Tissue glowing on its own can cause background issues.

Enzyme Methods: Use antibodies with enzymes like horseradish peroxidase (HRP) or alkaline phosphatase (AP). These make a colored spot at the protein site when mixed with a color-making substance.

Several advanced enzyme-based detection systems exist to boost strength and cut background:

PAP Method (Peroxidase Anti-Peroxidase): An indirect method with a third layer of rabbit antibody to peroxidase linked with peroxidase. This gives high strength.

Avidin-Biotin Complex (ABC) Method: A common method with a biotin-linked secondary antibody and an avidin-biotin peroxidase mix. It can have high background from natural biotin.

Labeled Streptavidin Biotin (LSAB) Method: Like ABC, but uses streptavidin instead of avidin. Streptavidin is neutral and has no sugar groups, cutting non-specific sticking. LSAB is often stronger than ABC.

Polymeric Methods (e.g., EnVision, ImmPRESS): Use enzyme-linked polymer reagents. Many enzyme molecules are attached to a secondary antibody via a polymer (like dextran). These give high strength, low background, and fewer steps.

Catalyzed Signal Amplification (CSA) Methods (e.g., CSA, CSA II): Use Tyramide Signal Amplification (TSA) to place many marked tyramide molecules at the protein site. This boosts the signal a lot. They’re great for tiny protein amounts and weak antibodies. CSA II is biotin-free, avoiding background from natural biotin.

Here’s a quick look at strength levels for some detection methods:

Picking a detection system that gives exact, clear staining with enough strength is super important.

Chromogens

For enzyme-based systems, a chromogen is used as a base for the enzyme. It makes a colored spot where the protein is.

The chromogen choice depends on more than just color. It includes staining accuracy, strength, and how it works with counterstains and sealing materials.

DAB (3,3′-diaminobenzidine tetrahydrochloride): The top choice for HRP-based systems. It makes a brown spot. DAB is liked for its fast reaction, exact spot, and fit with common counterstains and permanent sealing materials.

AP Red / Fast Red: Used with AP-based systems, making a red spot. AP Red is often used for skin where brown DAB might blend with melanin.

Different colored chromogens can be used in one tissue slice for multiplexing (showing two or more proteins at once). The time with the chromogen should be set right. Too long can cause wrong positives or too much background. Too short can lead to wrong negatives.

Counterstaining

Counterstaining is a key step after main staining. It gives background contrast and highlights cell parts, usually nuclei.

Common color counterstains include Fast red, Hematoxylin, methyl green, and toluidine blue. For glowing detection, DAPI and Hoechst are common glowing nuclear counterstains. Hematoxylin is often used after color detection.

The counterstain should stand out from the chromogen. The amount of counterstaining needs control so it doesn’t hide weak positive staining. Some chromogens and counterstains might not work with organic solvents.

Sealing the Stained Sample

After staining, the slide should be sealed to keep the sample and staining safe. This is done by adding a coverslip with a sealing solution (mountant). Mounting keeps the tissue slice and stain stable.

Mountants can be organic or water-based. If glowing detection was used, an anti-fade agent should be in the mountant to stop fading. Water-based mounting media are suggested if the chromogen or counterstain doesn’t work with organic solvents. After the mountant sets, the coverslip can be sealed with nail polish or a store-bought sealant.

Sample Visualization

After staining and sealing, the tissue slices are viewed under a microscope. Color detection uses a light microscope. Glowing detection needs a glowing microscope.

New microscope tech gives better pictures. Digital pathology and high-content screening let you capture images, measure IHC data, and analyze lots of samples fast. Experts check the staining based on the amount, pattern, and place of the colored or glowing signal.

Steps to Better IHC Staining / Tips for Success

Getting clear and steady IHC staining needs close care at every step, from getting tissue to checking results. Many problems can pop up, like weak staining, extra background, errors, or wrong results. Here are some key tips from experts to improve IHC:

Tissue Prep: Think hard about tissue prep, whether freezing or fixing with formalin and embedding in paraffin. Make sure slicing is smooth and even. Fixing right is a must; avoid too little or too much fixing and dead areas. Use high-quality, thin, flat slices dried on charged or APES-coated slides. Skip protein-based glues on charged slides.

Protein Unmasking: Do heat or enzyme unmasking when needed. Fixing can hide protein parts. Heat unmasking is usually better than enzyme unless the antibody maker says otherwise. Set the buffer type (acidic or alkaline like citrate, EDTA, or Tris), temperature, and time right. Include a control without unmasking to spot errors.

Blocking: Block natural enzymes (like peroxidase, phosphatase) and biotin to stop wrong positives or errors when using detection systems that rely on them. Block non-specific antibody sticking spots with serum or a general blocker before adding the main antibody. More blocking time might help.

Antibody Choice and Setup: Know your antibody by testing its work and pickiness, ideally with Western blot. Pick your main antibody carefully for strength and exactness. Always check the antibody info sheet for fit and update with new batches. Test different amounts of main and secondary antibodies to find the best mix for strong signal and low background. Watch for antibody mix-ups.

Detection System: Choose the right detection system based on how much target protein is there and how strong the signal needs to be. Marked main antibodies work for high protein amounts. Marked secondary antibodies or polymer/signal boost systems are better for low-to-medium proteins.

Chromogen Choice: Pick a chromogen for staining accuracy, strength, and fit with counterstains and sealing materials. DAB is a common pick for HRP-based systems due to its traits and fit.

Counterstain and Sealing: Counterstaining gives background contrast and highlights nuclei. Choose a counterstain that stands out from your chromogen, especially in multiplexing. Make sure chromogens, counterstains, and sealing materials (water vs. organic) work together. Use water-based sealing if needed to protect staining.

Controls: Controls are super important for checking results and sharing data confidently.

Positive controls (tissue known to have the protein) show the method works. Run them on the same slide if possible.

Negative controls (like tissue without protein, no-main antibody control, or isotype control) prove staining is exact. Knock-out or knock-down tissue samples are great negative controls.

Internal positive and negative controls in the sample also help with quality checks.

Washing Steps: Use steady washing steps (time, amount, shaking) throughout to keep things even and remove loose reagents. Different washing ways can cause uneven background.

Avoiding Errors: Avoid problems like tissue drying, air bubbles, incomplete drying, old or badly stored reagents, and too-thick tissue slices. These can cause wrong results. Avoid uneven reagent spread by applying carefully.

Pre-Staining Standards: Prep starts when tissue is taken. Setting steady rules for tissue collection, fixing, processing, and slicing is key to keep shape and protein activity. Problems at these early steps can hurt IHC quality. Linking these steps and watching conditions adds big value to quality checks.

Check Results Carefully: Know what exact staining pattern and place to look for in your test slices and controls.

Multiple Labeling

Staining for two or more proteins at once in one tissue slice is often helpful. This can use different glowing dyes in glowing tests or enzyme-linked antibodies with different color-making substances that give unique colors. Careful planning is needed, including antibody amounts, conditions, and good color mixes. Basic ways include side-by-side, one-after-another, and next-to staining.

Immuno-Electron Microscopy

For super close-up views, IHC methods can be used for electron microscopy (EM). These are split into pre-embedding, where staining happens before resin setting, and post-embedding, where labeling is done after setting. Colloidal gold bits are often used as markers for immuno-EM. They’re easy to see under the electron microscope. Gold bits can be different sizes, allowing multiple staining at the EM level. Testing the main antibody’s traits and amount at the light microscope level is suggested before doing immuno-EM labeling.

Many companies offer reagents and solutions for good IHC staining. Celnovte also provide IHC solutions.

Celnovte Biotechnology provides a market-leading portfolio of IHC and ISH reagents and instruments. They feature proprietary intellectual property such as MicroStacker™.

Celnovte Biotechnology was founded in 2010 and has development and manufacturing facilities primarily located in China, with R&D subsidiaries in Shenzhen, Suzhou, and Maryland, USA. Their facilities in China are NMPA & GMP compliant and hold ISO13485 and ISO9001 certifications. Their self-cloned primary antibodies have received optimal ratings in NordiQC assessments for six consecutive years.

FAQ

Q: What is antigen retrieval and why is it needed?
A: Antigen retrieval (AR) is a prep step in IHC, especially for FFPE tissues. It unmasks hidden protein spots covered by fixation links. Fixation like formalin makes protein links that block antibody access to its target. AR methods, like heat-induced epitope retrieval (HIER) or proteolytic-induced epitope retrieval (PIER), break these links. This lets the main antibody bind exactly, boosting staining strength.

Q: Why are blocking steps important in IHC?
A: Blocking steps stop non-specific sticking and wrong positive staining. Blocking natural enzymes (like peroxidase or alkaline phosphatase) or biotin keeps these tissue parts from reacting with detection reagents and causing background. Blocking non-specific protein sticking spots on the tissue (e.g., with serum or a buffer) stops antibodies from sticking to wrong places, cutting background staining.

Q: What’s the difference between direct and indirect IHC methods?
A: The difference is how the marker (for seeing) is attached to the antibody that binds the target. In the direct method, the main antibody is marked (e.g., with a glowing dye or enzyme) and binds the protein directly. This is one step but not very strong. In the indirect method, an unmarked main antibody binds the protein. Then a marked secondary antibody binds the main antibody. This is stronger because many marked secondary antibodies can bind one main antibody, boosting the signal.

Q: Why are controls necessary in IHC tests?
A: Controls are super important for checking if IHC staining results are reliable and exact. Positive controls (tissue known to have the target protein) show the method works right and reagents are good. Negative controls (like skipping the main antibody or using an isotype control) prove the staining is specific to the main antibody and not from non-specific sticking or errors. Without controls, you can’t trust the results or tell true staining from background or wrong signals.

Q: What are some common problems that can hurt IHC quality during sample prep?
A: Problems during sample prep can really affect IHC results. These include tissue damage during removal, lack of oxygen from delays before fixing, wrong sample labels, and uneven or wrong fixation (too little, too much, or irregular). Issues during slicing (too thick, tissue harm, cross-mixing) and processing (incomplete or over-fixation, bad decalcification, wrong schedule, poor reagents) also hurt quality. Embedding problems like wrong positioning or overloaded holders, and slicing issues like bad knives or rushed work, can cause errors and affect staining evenness and quality.