IHC on Frozen Tissue

Our protocol for performing IHC on frozen tissue sections.

Preparing Snap-Frozen Tissues for Post-Sectioning Fixation

Here we describe how to snap-freeze tissue, section tissue on a cryostat, and then postfix on a slide

• Place the freshly-dissected tissue block (<5mm thick) onto a mold.
• Use cryo-embedding media (e.g. OCT) to fully cover the entire tissue block.
• Completely freeze the tissue on the base mold by submersion in liquid nitrogen.
• Store the frozen tissue block at -80°C until ready for sectioning (See section on frozen sectioning below).
• Once sectioned, fix slides by incubating them in either ice-cold acetone (-20°C) for 20 minutes or 4% PFA

Preparing Fixed Tissues for Frozen Sectioning

Below is a general protocol for fixing tissues before sectioning on the cryostat.

• When possible, fixed perfusion is recommended for achieving the best tissue morphology and clear background. Perfuse the tissue with ice-cold PBS to remove blood, followed by fixation with 4% PFA until adequate fixation is achieved. Alternatively, dissect the desired tissues and fix them in 4% PFA or 10% formalin overnight (but no longer than 24 hours). The adequate volume of fixative for fixation by immersion is 50-100x of the sample size.
• Tissues should then be dehydrated for cryoprotection in fresh 20-30% sucrose, for 16-48 hours at 4°C.
• Embed the tissues in OCT medium using a mold and freeze by submersion in liquid nitrogen.
• Store the tissues at -80°C until ready for sectioning.

Sectioning Tissues on a Cryostat

• Optimal cryostat temperatures recommended for different tissue types are displayed in the table below.
• Before sectioning, transfer the frozen tissue block to a cryostat at the recommended temperature (e.g. -20°C) and allow the frozen tissue block to equilibrate to the cryostat temperature (around 30 mins).
• Section the frozen tissue block into a desired thickness. Use glass slides (we recommend charged slides) designed for IHC to prevent tissue slippage.
• Once cut, allow sections to dry thoroughly at room temperature. Sections can be stored in a sealed slide box at -80°C for later use. Storage might influence the antigenic potential, which varies for each protein. Therefore, minimizing storage time or using freshly prepared slides prior to staining is recommended.
• If necessary, slides can be fixed immediately after being removed from the freezer with either ice-cold acetone (-20°C) or 4% PFA for 10-20 minutes. The end user must determine the optimal fixation conditions for each tissue type.

Tissue Type	Temperature
Brain, liver, and lymph node tissues	-10°C/-15°C
Thyroid, spleen, kidney, and mouse tissues	-15°C/-20°C
Fat-containing tissue	-25°C
Tissue containing plenty of fat	-30°C

Antigen Retrieval (Optional)

• Please note that antigen retrieval is not required for frozen tissues if they were not fixed or if fixation was done with an alcohol, as these solutions do not mask epitopes.
• Transfer slides to a microwave-proof container or a beaker on an electric stove and cover them with the recommended antigen retrieval buffer. If no antigen retrieval buffer is suggested, try TRIS-EDTA (pH 9) first. See “IHC Antigen Retrieval” section for more information.
• Heat in the microwave for 10 minutes on medium power.
• Allow slides to cool in the antigen retrieval buffer for approximately 35 minutes.

Quenching and Blocking

• Rinse slides three times with 1x TBS for 3 minutes each.
• Incubate slides with 3% H2O2. (diluted in distilled water) for 10 minutes to quench endogenous peroxidase activity. For FFPE samples, this step is often not necessary. See the section on “Quenching” for more information.
• Rinse slides three times with 1x TBS for 3 minutes each.
• Prepare 1% BSA or 5% normal blocking serum in 1x TBS. The serum should be derived from the same species in which the secondary antibody was raised. Block the sections for 30-60 mins.

Primary Antibody Incubation

• Incubate sections with primary antibody diluted in 1x TBS for 1 hour, or overnight at 4°C; the optimal antibody dilution ratio should be pre-determined by experimentation. Set up negative controls by omitting the primary antibody incubation step for one slide per experimental condition.
• Following primary antibody incubation, rinse slides three times with 1x TBS for 3 minutes each.

Chromogenic Signal Detection

• Apply significant horseradish peroxidase (HRP) labeled secondary antibody (we recommend a Polymer-HRP conjugate for best results) and incubate for 30 minutes at room temperature.
• Rinse slides three times with 1x TBS for 3 minutes each.
• Prepare an appropriate volume of the DAB chromogen solution based on manufacturer guidelines. Apply the substrate carefully and incubate for 5-10 minutes until a brown color develops.
• Rinse sections gently with distilled water.
• Signal enhancement (optional): Immerse slides in 0.5-1% CuSO4 for 5 minutes.

Hematoxylin Counterstaining (optional)

• To stain nuclei, add a few drops of hematoxylin and incubate for 3 minutes.
• Rinse slides gently with distilled water.
• Transfer slides into a 1% HCl and 99% ethanol solution for 10 seconds and then rinse briefly in distilled water.

Fluorescent Signal Detection

• Add sufficient fluorophore labeled secondary and incubate at room temperature for 30 minutes in the dark. Ensure all steps following this are done in the dark or with minimal exposure to light.
• Optional: For counterstaining, we recommend DAPI to stain nuclei. This can be added here or use mounting media with DAPI.

Mounting

• Mount the sections with sufficient mounting media and cover with a cover slip. Air-dry in a well-ventilated area (e.g. fume hood).