Immunohistochemistry (IHC) fixation protocol

​

Fixation methods for cell culture samples

The correct fixative to use must be carefully considered for each IHC experiment as inappropriately fixed antigens may not be detected. 

• Formalin
• Ethanol
• Methanol
• Acetone

Read our in-depth guidelines on IHC fixation here

Formalin

• Add 10% neutral buffered formalin (NPF) to slides for 10 min.
• Wash 3x with PBS.

Fixing in formalin for more than 10–15 min will cross-link the proteins to the point where antigen retrieval may be required to ensure the antibody has free access to bind and detect the protein.

Ethanol

• Add 100–200 μL of 95% ethanol, 5% glacial acetic acid per slide.
• Place at -20°C for 5–10 min preferably in Coplins jar.
• Wash 3x with PBS.

Methanol

• Add 100–200 μL of ice-cold methanol, 5% acetic acid per slide.
• Place at -20°C for 10 min preferably in Coplins jar
• Wash 3x with PBS.

Ethanol and methanol will also permeabilize. Some epitopes are very sensitive to methanol as it can disrupt epitope structure. If this is occurring, try acetone instead if permeabilization is required.

Acetone

• Add 100–200 μL ice cold acetone per slide.
• Place at -20°C for 5-10 min.
• Wash 3x with PBS.

Acetone will also permeabilize. No further permeabilization step is required. 

Fixation methods for tissue samples

• Immersion fixation
• Perfusion fixation

Immersion fixation

10% neutral buffered formalin (NBF) is most commonly used. Where our datasheets state IHC-P as a tested application, this fixative has been used unless stated otherwise. Other fixatives such as Bouin solution (paraformaldehyde/picric acid) are used less frequently.

The ideal fixation time will depend on the size of the tissue block and the type of tissue, but fixation between 18–24 h seems to be suitable for most applications. Under-fixation can lead to edge staining, with 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, but if the tissue has been fixed for a long period of time (i.e. over a weekend), there may be no signal even after antigen retrieval.

Perfusion fixation

Perfusion fixation involves dissecting an animal and flushing 4% paraformaldehyde through its circulatory system via the heart. The tissue of interest can then be extracted and fixated further with immersion in the fixative.

Frozen section fixation

1. Once mounted on 3-amino-propyl-tri-ethoxy-silane (APES)-coated slides, sections should be air dried under airflow for 30–60 min to ensure complete desiccation as possible.
2. Store sections at -80°C until needed. For best results, use immediately.
3. When required, leave to warm at room temperature for 5 min.
4. Prepare fixative (acetone, methanol or ethanol) at room temperature. For a new antibody, we recommend starting with three sides:
   
   1) Paraformaldehyde
   2) Acetone
   3) 1:1 solution of acetone:alcohol (methanol or ethanol)
5. Fix with the fixative for 15 min, at room temperature.
6. Rinse 3–4 times in PBS.
7. For acetone fixation, air dry completely for 30 min under airflow.
8. Continue with the immunohistochemical staining protocol.

If the tissue samples are fixed with an aldehyde fixative (paraformaldehyde, glutaraldehyde etc) for immunofluorescence detection, include 0.3 M glycine in the blocking buffer, before applying the primary antibody.

Glycine will bind free aldehyde groups that would otherwise bind the primary and secondary antibodies, leading to high background. Background due to free aldehyde groups is more likely to occur when the fixative is glutaraldehyde or paraformaldehyde.