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Note: These protocols are provided on an ‘as is’ basis and may require optimization for your experimental conditions.
There are two methods for fixing tissues before embedding:
Adequate fixation is usually obtained by immersing small tissue pieces into the fixative solution. Still, perfusion can result in a more rapid and uniform fixation that preserves tissues in a life-like state.
Cut out a block of tissue from the source and wash it in PBS.
Use clean tools and keep samples on ice.
Fix the sample by incubating it with your chosen fixative.
For detection of | Suggested fixative |
Most proteins, peptides and enzymes of low molecular weight. | 10% Neutral-Buffered Formalin (NBF) or 4% paraformaldehyde (PFA)* |
Small molecules, such as amino acids | 10% Neutral-Buffered Formalin (NBF) or 4% paraformaldehyde (PFA)* |
Large protein antigens, such as immunoglobulin | 100 % acetone or methanol |
Table 1. Common fixatives in IHC
*Note that the concentrations of formaldehyde in both 10% NBF and 4% PFA are almost identical.
10% neutral buffered formalin (NBF) is the most commonly used.
Where our datasheets state IHC-P as a tested application, NBF fixative has been used unless stated otherwise.
Note that acetone and methanol will also permeabilize samples.
The fixation time may require optimization: under-fixation can lead to edge staining, with a strong signal on the edges of the section and no signal in the middle. Over-fixation can mask the epitope; antigen retrieval can help overcome this masking.
Wash the tissue three times with PBS.
Following fixation, samples are now ready to be embedded in paraffin. This procedure can be carried out using a vacuum oven to heat the paraffin or using an automated embedding system.
Rinse the tissue with PBS until the fixative is completely removed.
Dehydrate the tissue at room temperature with ethanol, then clear it with xylene.
Solution | Incubation time |
50% Ethanol | 1-2 hrs |
75% Ethanol | 20-30 mins |
85% Ethano | 20-30 mins |
85% Ethanol | 20-30 mins |
95% Ethanol | 20-30 mins |
95% Ethanol | 20-30 mins |
100% Ethanol | 20-30 mins |
100% Ethanol | 20-30 mins |
Xylene | 10-20 mins |
Xylene | 10-20 mins |
Some larger pieces of tissue can benefit from slower dehydration, i.e. longer incubation times.
Tissue processing cassettes are very useful for containing samples and are then embedded at the back of your wax block to be used as an anchoring point for sectioning.
You can use several less hazardous commercially available alternatives to xylene. Please refer to the manufacturer’s instructions for ethanol exchange and mounting procedures specific to the agent.
Exchange the tissue three times in hot paraffin.
Solution | Incubation time |
Paraffin | 40 mins |
Paraffin | 1 hr |
Paraffin | 1 hr |
Various manufacturers supply paraffin wax. Be careful to use the correct oven temperature for the wax you are using, and refer to the manufacturer if unsure.
Embed the tissue mold in fresh paraffin.
Once the tissue is embedded, we can cut it into sections and mount it to microscope slides.
Chill paraffin-embedded tissue blocks on ice.
Cold wax allows thinner sections to be obtained by supporting harder elements within the tissue specimen.
The small amount of moisture penetrating the block from the melting ice will make the tissue easier to cut.
Fill a water bath with ultrapure water and heat to 40 – 45°C.
Set up the microtome and blade.
The blade clearance angle should be adjusted to achieve optimum performance.
Insert the paraffin block into the microtome and orient.
Before cutting the block, carefully, you can cut a few thin sections to ensure the positioning is correct. Adjust if necessary.
Trim the block to expose the tissue surface.
Cut sections to a thickness of 3 – 10 µm.
You will probably need to discard the first few sections as they likely contain holes caused by trimming.
Separate the sections on a water bath and place them on microscope slides.
Dry the sections in an oven.
Drying the sections at 37°C overnight is less likely to damage heat-sensitive antigens.
Samples need to undergo deparaffinization to remove the wax and enable antibody penetration. Antigen retrieval steps will also help further expose the antigen and are often required for tissues fixed in formalin.
Before proceeding to antibody incubation, we must deparaffinize and rehydrate the slides. Incomplete removal of paraffin can lead to poor staining of the section.
Wash the slides in xylene and then in ethanol using a Coplin’s jar or autostainer.
Solution | Incubation time |
Xylene | 10-15 mins |
Xylene | 10-15 mins |
100% Ethanol | 5 mins |
100% Ethanol | 5 mins |
95% Ethanol | 5 mins |
95% Ethanol | 5 mins |
85% Ethanol | 5 mins |
85 % Ethanol | 5 mins |
75% Ethanol | 5 mins |
75% Ethanol | 5 mins |
These steps can be automated or manual.
You can use several less hazardous commercially available alternatives to xylene. Please refer to the manufacturer’s instructions for exchanging ethanol and mounting procedures specific to the agent.
Wash the slides three times with water.
Keep the slides in water until ready to perform antigen retrieval.
Most formalin-fixed tissues require an antigen retrieval step before immunohistochemical staining. Methylene bridges formed during fixation cross-link proteins and mask antigenic sites.
Antigen retrieval methods break these methylene bridges and expose antigenic sites, allowing antibodies to bind.
Immerse the slide rack into a staining dish containing antigen retrieval buffer.
Transfer the staining dish into a heating apparatus and heat the slides.
Equipment | Suggested method |
Decloaking chamber | 110 °C for 15 – 30 mins |
Scientific Microwave | 98 °C for 20 mins |
Slides should be placed in a plastic or metal rack and vessel. Standard glass histology staining racks and vessels will crack when heated.
If using a scientific microwave, set the temperature to a constant 98°C to avoid section dissociation from vigorous boiling.
Remove the slides from the vessel and run under cold tap water for 10 mins.
This allows the slides to cool enough so they may be handled, and allows the antigenic site to re-form after being exposed to high temperature.
Transfer slides back into the rack and immerse in water.
It's important to prevent the slides from drying out.
Now that we prepared the slides and antigen retrieval has taken place where required, the samples are ready for blocking and antibody incubation.
Blocking steps are particularly important in IHC to prevent high background staining in images. Blocking of endogenous proteins is essential for all samples.
Wash the slides twice with TBST.
Perform endogenous avidin/biotin block (optional).
Perform this step only for biotinylated antibodies.
Perform endogenous peroxidase block (optional).
Perform this step only for peroxidase-conjugated antibodies.
Perform protein block.
Perform this step only for all IHC experiments.
If using serum for blocking, the serum should match the host species of the secondary antibody.
Wash slides three times with TBST.
Proceed to immunostaining.
After performing the necessary blocking steps, we’re now ready to stain our tissue samples with antibodies. We can stain tissues directly with conjugated primary antibodies, or indirectly with conjugated secondary antibodies.
Multicolor IHC involves staining with two or more sets of antibodies to reveal the distribution of two or more proteins of interest. Both the indirect and direct protocols given below can be adapted for multicolor IHC. We can either:
Dilute the antibodies in TBST.
If not, you may need to perform dilutions to find the antibody concentration that works best.
1% BSA is included as an additional blocking reagent to reduce non-specific binding caused by hydrophobic interactions.
Incubate the slides with pre-diluted primary antibody.
Incubation time may need optimization.
The antibody solution needs to cover your samples completely.
Using a hydrophobic barrier pen can help contain small volumes.
Wash the slides three times with TBST.
Incubate the slides in pre-diluted secondary antibody.
Incubation time may need to be optimized.
If using a fluorophore, incubation must be in the dark to avoid photobleaching.
Endogenous peroxidase blocking can also be carried out at this point.
Wash the slides three times with TBST.
Proceed to counterstaining, mounting and imaging.
After incubation with antibodies, you’re now ready to image your slides according to the procedures below.
Immerse the slide in the fluorescent counterstain solution (optional).
See further advice on choosing a counterstain in our IHC guide.
Some mounting media are fortified with a fluorescent counterstain (see ab104139). This eliminates the need for an additional counterstain step.
Wash the slides with cold running water to remove the excess stain.
Add a few drops of mounting medium suitable for fluorescent detection to the slides.
Let the slides stand at room temperature for around 5 mins.
Use the minimum volume needed to mount the slides.
Carefully place a coverslip over the slide using forceps.
Be careful not to introduce any bubbles or disturb the sample.
Image the slides using a microscope.
If not using immediately, store slides in the dark at 4°C.