IHC with frozen samples
Protocol for performing IHC on frozen samples
Immunohistochemistry (IHC) on frozen tissue sections is a widely used technique for detecting and localizing specific antigens within preserved cellular architecture. Unlike formalin-fixed paraffin-embedded (FFPE) samples, frozen tissues retain native antigenicity and are particularly suitable for targets sensitive to chemical fixation. This approach is essential for studying labile epitopes, post-translational modifications, and certain membrane proteins that may be masked or degraded during fixation.
Frozen IHC offers rapid sample processing and is commonly employed in translational research, neurobiology, and clinical diagnostics. However, it presents unique challenges, including tissue fragility, ice crystal artifacts, and variability in antigen preservation. Optimal results depend on careful sample handling, cryosectioning technique, and appropriate antibody validation.
This protocol outlines a reliable workflow for performing IHC on cryosections, from tissue preparation and fixation to antibody incubation and signal detection. It emphasizes critical steps such as fixation choice (e.g., acetone or paraformaldehyde), blocking strategies to reduce background, and compatibility with chromogenic or fluorescent detection systems. The protocol is adaptable to a range of tissue types and target proteins, and includes troubleshooting guidance to improve reproducibility and signal specificity.
By following this protocol, researchers can achieve high-quality staining with minimal background, enabling accurate interpretation of protein localization and expression in frozen samples.
Stage 1 - Freezing
To prepare tissue sections on slides, we must first freeze the samples before sectioning and fixation.
Materials required
- Samples
- Isopentane
- Dry ice
- Optimal Cutting Temperature (OCT) compound
- Tissue embedding molds and cassettes
Steps
Prepare a cold isopentane bath.
- Fill a large insulated container with dry ice.
- Partially fill a metal container with isopentane and place the container on dry ice.
- Wait for 5 mins, allowing the isopentane to be cooled by the dry ice.
The level of dry ice and isopentane should be the same.
The isopentane may freeze around the edges or at the bottom of the container; do not let the isopentane freeze completely.
Place fresh tissue in an embedding mold and fill it with OCT compound.
- Orient the tissue as desired
Freeze the tissue using the isopentane bath.
- Hold the tissue mold over the isopentane bath with forceps for 10 – 20 seconds until the tissue block turns opaque. This rapid freezing process is known as snap freezing, or to snap freeze the tissue, which helps preserve tissue morphology and integrity.
Store the frozen tissue at -80°C until ready for sectioning.
Stage 2 - Sectioning
Once the tissue is embedded, we can cut it into sections and mount it to microscope slides.
Materials required
- Frozen tissue samples
- Suitable slides
- Cryostat
- Cryostat blade
Steps
Bring frozen tissue samples to -20°C in a cryostat and allow them to equilibrate overnight.
Attach the frozen block to the sample holder and leave it to set.
Set up the cryostat blade by placing it in the holder, ensuring it is secure, and setting the clearance angle.
- Follow the manufacturer’s instructions for guidance on setting the clearance angle.
Trim the frozen tissue block to expose the tissue surface.
Proceed to cut sections at a thickness of 5 – 8 µm.
Collect the sections using a brush and place them on a slide ready for subsequent fixation steps.
Stage 3 - Fixation
Here we need to air dry the frozen samples and then fix them to preserve protein and tissue morphology before antibody incubation. Common fixatives include 10% neutral buffered formalin (NBF) and 4% paraformaldehyde (PFA), both of which are examples of formaldehyde based fixation used to ensure tissue integrity and antigen preservation.
Materials required
- Tissue sections on suitable IHC slides
- Wash buffer (PBST)
- Fixative (10% NBF or 4% PFA, 100% acetone or methanol)
Steps
Dry frozen samples at room temperature for 15 mins.
Select a suitable fixative at room temperature.
Most proteins, peptides, and enzymes of low molecular weight
Small molecules like amino acids
For a new antibody, we recommend starting with three sides:
1) 4% Paraformaldehyde
2) 100% methanol
3) 1:1 solution of acetone:alcohol (methanol or ethanol)
Note that the concentrations of formaldehyde in both 10% NBF and 4% PFA are almost identical.
Immerse the tissue slide in the fixative solution.
- Incubate samples for 15 mins at room temperature.
Wash the tissue slides three times with PBST.
Stage 4 - Blocking
Blocking steps are particularly important in IHC to prevent high background staining in images. Using a blocking buffer is essential to reduce non-specific binding of antibodies, thereby improving staining specificity and clarity. It is important to note that the blocking buffer is applied before antibody incubation, and is distinct from the incubation buffer used during the primary and secondary antibody incubation steps.
Materials required
- Your tissue sections
- 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)
- Endogenous peroxidase blocking solution (optional for peroxidase-conjugated antibodies – example: ab64218)
- Wash buffer, PBST (PBS, 0.05% Triton X-100 or NP-40)
Steps
Wash the slides twice for 5 mins in PBST.
Perform endogenous avidin/biotin block (optional).
- Incubate slides for 10 mins in avidin blocking solution at room temperature.
- Wash slides once with PBST.
- Incubate slides for 10 mins in biotin-blocking solution at room temperature.
- Wash slides once with PBST.
Perform endogenous peroxidase block (optional).
- Incubate slides with 3% hydrogen peroxide for 10 mins at room temperature.
- Wash slides once with PBST.
Perform protein block.
- Incubate slides in protein blocking reagent for 30 – 60 mins at room temperature.
Wash slides three times for 5 mins with PBST.
Proceed to immunostaining.
Stage 5 - Antibody incubation
After performing the necessary blocking steps, we’re ready to stain our tissues with antibodies. We can stain tissues directly with conjugated primary antibodies or indirectly with conjugated secondary antibodies.
Multicolor IHC involves staining cells with two or more 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, or incubate cells with each antibody set sequentially, performing blocking between each incubation.
Materials required
- Tissues that have undergone relevant blocking steps
- Antibody dilution buffer (PBS + 1% BSA)
- Wash buffer (PBST)
- Conjugated primary antibody
Steps
Determine the optimal antibody dilutions to use, then dilute the antibodies in PBS 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.
Incubate the slides in the pre-diluted primary antibody.
- Incubate for 1 hr at room temperature, or 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 PBST.
Proceed to counterstaining, mounting and imaging.
Materials required
- Tissues that have undergone relevant blocking steps
- Dilution buffer (PBS, 1% BSA)
- Wash buffer (PBST)
- Primary antibody
- Conjugated secondary antibody (example: our conjugated secondaries)
- Hydrophobic barrier pen (example: ab2601)- optional
Steps
Dilute the primary and secondary antibodies in PBS with 1% BSA.
- Suggested dilutions will often be suggested on the antibody datasheet.
Incubate the samples with the pre-diluted primary antibody.
- Incubate for 1 – 2 hrs at room temperature, or 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 PBST.
Incubate the samples with pre-diluted secondary antibody.
- Incubate according to the manufacturer’s guidance.
- Usually, it’s recommended to incubate for 45-60 mins at room temperature.
Incubation time may need to be optimized.
If using a fluorophore, incubation must be in the dark to avoid photobleaching.
Before adding an HRP-conjugated secondary antibody, you can carry out endogenous peroxidase blocking at this point.
Wash the slides three times with PBST.
Stage 6 - Detection
After incubation with antibodies, you're now ready to image your antibodies according to the procedures below.
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
Immerse the slide in chromogenic substrate solution.
- Incubate until the desired color is observed.
Monitor the staining visually during the incubation.
AP blocking can be included here if required.
Wash the slides with cold running water to remove 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 and clear the tissue before applying organic mounting media (optional).
- Perform the following exchange at room temperature, manually in a Coplin’s jar, or in an automated embedding system.
Add a few drops of mounting medium to the slid: let the slide stand at room temperature for 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.
-
If using an organic mounting medium, allow the medium to dry completely.
Image the slides using a microscope.
Materials required
- Tissue slides stained with fluorophore-conjugated antibody
- Fluorescent counterstain (optional, example: DAPI ab228549)
- Mounting medium suitable for fluorescent detection (example: ab104135)
- Sealant (example: nail polish or Limonene ab104141)
- Coverslip
- Microscope
- Deionized (DI) water
Steps
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 DI water to remove the excess stain.
Add a few drops of mounting medium to the slides.
- Let slide 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.
Imaging lab essentials
- Marker antibodies
- Immunostaining, detection systems and counterstains
- Isotype controls
- Buffers, mounting media and other accessories
Easier IHC with validated antibodies for BOND RX
- BOND RX kitted antibodies
- BOND RX validated antibodies
- Enhanced validated antibodies
- mIHC antibodies