IHC with samples in paraffin (IHC-P)
By our in-house imaging experts
Note: These protocols are provided on an ‘as is’ basis and may require optimization for your experimental conditions.
Stage 1 - Sample preparation: fixation
There are two methods for fixing tissues before embedding:
- Immersion fixation for when the tissue has already been excised
- Perfusion for fixation in situ through the vascular system (here, we share a couple of example protocols)
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.
Materials required
- Samples
- PBS
- Fixative (example: 10% neutral buffered formalin (NBF)
Steps
Cut out a block of tissue from the source and wash it in PBS.
Fix the sample by incubating it with your chosen fixative.
- Immerse the tissue block in one of the following solutions for 18 – 24 hrs at 4°C.
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.
Perfusion involves dissecting an anesthetized animal and flushing 4% paraformaldehyde (PFA) through its circulatory system via the heart.
After this process, the tissue of interest is extracted, and further fixation via immersion can be performed.
For more detailed information on perfusion, you can refer to the following resources:
· Whole animal perfusion fixation for rodents (for brain tissue preservation: Gage, G., Kipke, D. and Shain, W., 2012. Whole Animal Perfusion Fixation for Rodents. Journal of Visualized Experiments, (65).
· Perfusion fixation through the heart and abdominal aorta in rats: Celis, J., 2006. Cell biology. 3rd ed. London: Academic Press, pp.225-227.
Stage 2 - Sample preparation: embedding
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.
Materials required
- Samples
- PBS
- Ethanol
- Xylene or another clearing agent
- Paraffin wax
- Tissue embedding molds and cassettes
- Vacuum oven or automated embedding system
Steps
Rinse the tissue with PBS until the fixative is completely removed.
Dehydrate the tissue at room temperature with ethanol, then clear it with xylene.
- Perform the following exchange at room temperature, either manually in a Coplin’s jar or in an automated embedding system.
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.
- Check the paraffin manufacturer’s instructions for recommended temperature – typically 50 – 60°C.
- Perform the exchanges in a vacuum oven or an automated embedding system.
Embed the tissue mold in fresh paraffin.
- Carefully place the tissue in a mold and fill the mold with paraffin.
- Orient the tissue as desired and use a cassette to anchor the tissue to the mold.
- Place the tissue block in the fridge for ~10 mins to harden the paraffin.
Stage 3 - Sample preparation: sectioning
Once the tissue is embedded, we can cut it into sections and mount it to microscope slides.
Materials required
- Samples
- Waterbath
- Container with ice
- Glass microscope slides
- Microtome and blade
- Oven or slide drying rack
Steps
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.
- Place the blade in the holder.
- Follow the microtome manufacturer’s instructions for guidance on setting the clearance angle.
Insert the paraffin block into the microtome and orient.
Trim the block to expose the tissue surface.
- Trimming is normally done to a thickness of 10 – 30 µm.
- This allows more representative sections to be cut later.
Cut sections to a thickness of 3 – 10 µm.
- Sections will come off as ribbons.
Separate the sections on a water bath and place them on microscope slides.
- Pick up the ribbons of sections and float them on the surface of the water bath, so they flatten out.
- Use tweezers to separate the sections.
- Pick the sections from the water bath onto a microscope slide and place them in a slide rack.
Dry the sections in an oven.
Stage 4 - Deparaffinization
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.
Materials required
- Samples
- Xylene
- Ethanol
- Coplin Jars or autostainer (example: Leica ST5020 Multistainer)
Steps
Wash the slides in xylene and then in ethanol using a Coplin’s jar or autostainer.
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.
Stage 5 - 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.
In heat-induced epitope retrieval (HIER), we place the tissue sections in a buffer and heat them to expose antigenic sites.
HIER is most often performed using a water bath, pressure cooker, or scientific microwave.
Materials required
- Suitable antigen retrieval buffer (example: Tris EDTA buffer, ab93684)
- Your tissue sections, deparaffinized and hydrated in water
- Slide rack to hold approximately 400–500 mL
- Heating apparatus (decloaking chamber or scientific microwave)
Steps
Immerse the slide rack into a staining dish containing antigen retrieval buffer.
Transfer the staining dish into a heating apparatus and heat the slides.
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.
Transfer slides back into the rack and immerse in water.
In enzymatic antigen retrieval, the tissue sections are incubated with a protease such as Trypsin, Pepsin, or Proteinase K to expose the antigenic sites. The enzyme can be applied to slides directly with a pipette, or the slide rack can be immersed into the enzyme solution.
Materials required
- Your tissue sections on glass slides, deparaffinized and hydrated in water
- Slide rack
- The incubator set to the optimum temperature (usually 37°C)
- Enzymatic antigen retrieval solution (examples: Trypsin ab970, Pepsin ab64201 Proteinase K ab64220)
Steps
Prepare the enzymatic antigen retrieval solution as recommended by the manufacturer.
The best enzyme to use should be indicated on the antibody datasheet. If not, you can use Trypsin to retrieve various antigens after formalin fixation.
Our enzymatic antigen retrieval solutions are ready to use. However, check the literature if using another manufacturer; some enzymes require specific buffers and cofactors for activity.
Heat the enzymatic antigen retrieval solution in an incubator to the optimum temperature for the enzyme you’re using.
- Trypsin: 37°C
- Pepsin: 37°C
- Proteinase K: room temperature
Treat tissue slides with the pre-heated enzymatic antigen retrieval solution.
This can be done by:
- Pipetting a small amount (50 – 100 µL) of the enzymatic solution onto the slides, or
- Immersing the slide rack into the solution, keeping slides immersed throughout step 4.
Place the treated slides into an incubator and heat for ~5 – 10 mins.
- Trypsin: 37°C
- Pepsin: 37°C
- Proteinase K: room temperature
If pipetting onto slides directly, keep the incubator humidified to prevent the slides from drying out. You can do it by using a humidified chamber.
Use a slide rack to avoid placing slides directly on the incubator shelves.
An incubation time of 5 – 10 mins will generally work, but you may wish to optimize by trying a range of times between 5 and 30 minutes.
Rinse slides under cold running water to remove excess enzyme.
Transfer slides into the rack and immerse in water.
Stage 6 - Blocking
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.
Materials required
- TBST (example: ab64202 or 1×TBS/0.1% Tween-20, pH 7.4±0.2)
- Protein blocking solution (example 2 - 10 % normal serum of same species as secondary antibody, or sera-free protein block ab64226)
- Biotin blocking solution (optional – for biotinylated antibodies – example ab64212)
- 3%vHydrogen peroxide (optional for peroxidase-conjugated antibodies – example ab64218)
Steps
Wash the slides twice with TBST.
Perform endogenous avidin/biotin block (optional).
- Incubate slides for 10 mins in avidin blocking solution at room temperature.
- Wash slides once with TBST.
- Incubate slides for 10 mins in biotin-blocking solution at room temperature.
- Wash slides once with TBST.
Perform endogenous peroxidase block (optional).
- Incubate slides with 3% hydrogen peroxide for 10 mins at room temperature.
- Wash slides once with TBST.
Perform protein block.
- Incubate slides in protein blocking reagent for 30 – 60 mins at room temperature.
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.
Stage 7 - Antibody incubation (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:
- Incubate cells with multiple antibody sets simultaneously.
- Incubate cells with each antibody set sequentially, performing blocking between each incubation.
Materials required
- Tissues that have undergone relevant blocking steps
- TBST (example: ab64248)
- BSA
- Primary antibody
- Conjugated secondary antibody
- Optional: Hydrophobic barrier pen (example: ab2601)
- Automated stainer
Steps
Dilute the antibodies in TBST.
- Optimum dilutions will often be suggested on the antibody datasheet.
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.
- Immerse slides in antibody solution overnight at 4°C.
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.
- Immerse slides in antibody solution for around 10-30 mins at room temperature. Please refer to the manufacturer’s guidance for exact incubation conditions.
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.
Materials required
- Tissues that have undergone relevant blocking steps
- TBS (example: ab64248)
- BSA
- Conjugated primary antibody
Steps
Dilute the antibodies in TBS with 1% BSA.
- Optimum dilutions will often be suggested on the antibody datasheet.
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 in the pre-diluted primary antibody.
- Incubate overnight at 4°C.
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 TBS.
Proceed to counterstaining, mounting and imaging.
Stage 8 - Detection
After incubation with antibodies, you’re now ready to image your slides according to the procedures below.
Materials required
- Tissue slides stained with fluorophore-conjugated antibody
- Fluorescent counterstain (optional, example: DAPI ab228549)
- Mounting medium suitable for fluorescent detection
- Sealant (example: nail polish or Limonene ab104141)
- Coverslip
- Microscope
Steps
Immerse the slide in the fluorescent counterstain solution (optional).
- Incubate according to the manufacturer’s guidance at room temperature or until the desired color is observed.
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.
Carefully place a coverslip over the slide using forceps.
- If using an aqueous mounting medium, seal the coverslip with limonene or nail polish.
Image the slides using a microscope.
Materials required
- Tissue slides stained with enzyme conjugated antibody
- Chromogenic substrate (examples for HRP: DAB ab64238, AEC ab64252)
- Counterstain (optional, example: Mayer’s Hematoxylin ab220365)
- Mounting medium (organic, example ab104141, or aqueous ab64230)
- Sealant (optional for aqueous mounting media, example nail polish or Limonene ab104141)
- Coverslip
- Microscope
Steps
Incubate the slides with chromogenic substrate.
- Immerse slides in the substrate solution at room temperature until the desired color is observed.
Freshly prepare your chromogenic substrate according to the product specification and choose suitable staing time and cycles.
Monitor the staining visually during the incubation.
AP blocking can be included here if required.
Wash the slides with cold running water to remove the excess stain.
Immerse the slide in counterstain solution (optional).
- Incubate according to the manufacturer’s guidance at room temperature or until the desired color is observed.
Wash the slides with cold running water to remove the excess stain and to blue the hematoxylin.
Dehydrate the tissue using ethanol and xylene.
- Perform the following exchange at room temperature, either manually in a Coplin’s jar or in an automated embedding system.
Add a few drops of mounting medium to the slides and let the slide stand at room temperature for 5 mins.
Place a coverslip over the slide using forceps.
- If using an aqueous mounting medium, seal the coverslip with limonene or nail polish.
- If using an organic mounting medium, allow the medium to dry completely.
Image the slides using a microscope.