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IHC with samples in paraffin (IHC-P)

By our in-house imaging experts
Last edited Wed 22 Feb 2023

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

1

Cut out a block of tissue from the source and wash it in PBS.

2

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.
For detection ofSuggested fixative
Most proteins, peptides and enzymes of low molecular weight.10% Neutral-Buffered Formalin (NBF) or 4% paraformaldehyde (PFA)*
Small molecules, such as amino acids10% Neutral-Buffered Formalin (NBF) or 4% paraformaldehyde (PFA)*
Large protein antigens, such as immunoglobulin100 % 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.

3

Wash the tissue three times with PBS.

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

1

Rinse the tissue with PBS until the fixative is completely removed.

2

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.
SolutionIncubation time

50% Ethanol

1-2 hrs 
75% Ethanol20-30 mins
85% Ethano20-30 mins
85% Ethanol20-30 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
Xylene10-20 mins
3

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.
SolutionIncubation time
Paraffin40 mins
Paraffin1 hr
Paraffin1 hr

 

4

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

1

Chill paraffin-embedded tissue blocks on ice.

2

Fill a water bath with ultrapure water and heat to 40 – 45°C.

3

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.

Figure 1. Blade clearance angle. Setting the right clearance angle prevents contact between the knife facet and the face of the block.

4

Insert the paraffin block into the microtome and orient.

5

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.
6

Cut sections to a thickness of 3 – 10 µm.

  • Sections will come off as ribbons.
7

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.
8

Dry the sections in an oven.

  • Place the slide rack into an oven at 63 – 65 °C for 30 – 60 mins or at 37°C overnight.

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

Steps

1

Wash the slides in xylene and then in ethanol using a Coplin’s jar or autostainer.

SolutionIncubation time
Xylene10-15 mins
Xylene10-15 mins
100% Ethanol5 mins
100% Ethanol     5 mins
95% Ethanol      5 mins
95% Ethanol5 mins
85% Ethanol5 mins
85 % Ethanol5 mins
75% Ethanol5 mins
75% Ethanol5 mins
2

Wash the slides three times with water.

3

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.

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

1

Immerse the slide rack into a staining dish containing antigen retrieval buffer.

2

Transfer the staining dish into a heating apparatus and heat the slides.

EquipmentSuggested method
Decloaking chamber110 °C for 15 – 30 mins
Scientific Microwave98 °C for 20 mins
3

Remove the slides from the vessel and run under cold tap water for 10 mins.

4

Transfer slides back 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

1

Wash the slides twice with TBST.

2

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.
3

Perform endogenous peroxidase block (optional).

  • Incubate slides with 3% hydrogen peroxide for 10 mins at room temperature.
  • Wash slides once with TBST.
4

Perform protein block.

  • Incubate slides in protein blocking reagent for 30 – 60 mins at room temperature. 
5

Wash slides three times with TBST.

6

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

1

Dilute the antibodies in TBST.

  • Optimum dilutions will often be suggested on the antibody datasheet.
2

Incubate the slides with pre-diluted primary antibody.

  • Immerse slides in antibody solution overnight at 4°C. 
3

Wash the slides three times with TBST.

4

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.
5

Wash the slides three times with TBST.

6

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

1

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.
2

Wash the slides with cold running water to remove the excess stain.

3

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.

4

Carefully place a coverslip over the slide using forceps.

  • If using an aqueous mounting medium, seal the coverslip with limonene or nail polish.
5

Image the slides using a microscope.