Immunohistochemical staining protocol for frozen samples (IHC-Fr)
Licia Miller Product Manager
Stage 1 Freezing
To prepare tissue sections on slides, we must first freeze the samples before sectioning and fixation.
Materials required
- Samples
- Dry ice
- Optimal Cutting Temperature (OCT) compound
- Tissue embedding molds and cassettes
Steps
1.Prepare a cold isopentane bath.
1.1 Fill a large insulated container with dry ice.
1.2 Partially fill a metal container with isopentane and place the container on dry ice.
1.3 Wait for 5 mins, allowing the isopentane to be cooled by the dry ice.
Tips: 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.
2.Place fresh tissue in an embedding mold and fill it with OCT compound. And orient the tissue as desired.
3.Freeze the tissue using the isopentane bath.
3.1 Hold the tissue mold over the isopentane bath with forceps for 10-20 seconds until the tissue block turns opaque.
Notice: Avoid getting any isopentane on the OCT compound.
4.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
1.Bring frozen tissue samples to -20°C in a cryostat and allow them to equilibrate overnight.
Notice: Ensure all tools used are kept in the cryostat and brought to temperature before cutting.
2.Attach the frozen block to the sample holder and leave it to set.
3.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.
Notice: The blade clearance angle should be adjusted to achieve optimum performance.
4.Trim the frozen tissue block to expose the tissue surface. Trimming is normally done at a thickness of 10-30 µm.
5.Proceed to cut sections at a thickness of 5-8 µm.
Tips: You will probably need to discard the first few sections as they likely contain holes caused by trimming.
6.Collect the sections using a brush and place them on a slide ready for subsequent fixation steps.
Tips: A toothpick can also be used, or the section can be lifted slightly, and the slide can be placed against the section, avoiding contact with the stage, which will then adhere to the slide.
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.
Materials required
- Tissue sections on suitable IHC slides
- Wash buffer (PBST)
- Fixative (10% NBF or 4% PFA , 100% acetone or methanol)
Steps
1.Dry frozen samples at room temperature for 15 mins.
Tips: This procedure begins after freezing, embedding, and sectioning frozen samples. This is unlike wax embedding, where fixation takes place before embedding.
2.Select a suitable fixative at room temperature.
Table 1. Common fixatives used in IHC.
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Notice: 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.
3.Immerse the tissue slide in the fixative solution. Incubate samples for 15 mins at room temperature.
4.Wash the tissue slides three times with PBST.
Notice: For acetone fixation, air dry completely for 30 min under airflow.
Stage 4 Blocking
Blocking steps are particularly important in IHC to prevent high background staining in images.
Materials required
- Your tissue sections
- Protein blocking solution (example: 2-10% normal serum of same species as secondary antibody, or sera-free protein block )
- Biotin blocking solution (optional – for biotinylated antibodies )
- Endogenous peroxidase blocking solution (optional)
- Wash buffer, PBST (PBS, 0.05% Triton X-100)
Steps
1.Wash the slides twice for 5 mins in PBST.
2.Perform endogenous avidin/biotin block (optional).
2.1 Incubate slides for 10 mins in avidin blocking solution at room temperature.
2.2 Wash slides once with PBST.
2.3 Incubate slides for 10 mins in biotin-blocking solution at room temperature.
2.4 Wash slides once with PBST.
Notice: You should perform this step only for biotinylated antibodies.
3.Perform endogenous peroxidase block (optional).
3.1 Incubate slides with 3% hydrogen peroxide for 10 mins at room temperature.
3.2 Wash slides once with PBST.
Notice: You should perform this step only for peroxidase-conjugated antibodies.
4.Perform protein block.
4.1 Incubate slides in protein blocking reagent for 30-60 mins at room temperature.
Notice: You should perform this step only for all IHC experiments.
If using a serum for blocking, the serum should match the host species of the secondary antibody.
The blocking solution should not contain serum of the host animal of the primary antibody as this will likely result in high background.
5.Wash slides three times for 5 mins with PBST.
6.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.
【Direct】
Materials required
- Tissues that have undergone relevant blocking steps
- Antibody dilution buffer (PBS+1% BSA)
- Wash buffer (PBST)
- Conjugated primary antibody
Steps
1.Determine the optimal antibody dilutions to use, then dilute the antibodies in PBS with 1% BSA.
Notice: 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.
2.Incubate the slides in the pre-diluted primary antibody for 1 hr at room temperature, or overnight at 4°C.
Notice: Incubation time may need optimization.
The antibody solution needs to cover your samples completely.
Using a hydrophobic barrier pen can help contain small volumes.
3.Wash the slides three times with PBST.
4.Proceed to counterstaining, mounting and imaging.
【Indirect】
Materials required
- Tissues that have undergone relevant blocking steps
- Dilution buffer (PBS, 1% BSA)
- Wash buffer (PBST)
- Primary antibody
- Conjugated secondary antibody
- Hydrophobic barrier pen- optional
Steps
1.Dilute the primary and secondary antibodies in PBS with 1% BSA.
Notice: Suggested dilutions will often be suggested on the antibody datasheet.
You may need to perform dilutions to find the antibody concentration that works best.
2.Incubate the samples with the pre-diluted primary antibody for 1-2 hrs at room temperature, or overnight at 4°C.
Notice: Incubation time may need optimization.
The antibody solution needs to cover your samples completely.
Using a hydrophobic barrier pen can help contain small volumes.
3.Wash the slides three times with PBST.
4.Incubate the samples with pre-diluted secondary antibody according to the manufacturer’s guidance. Usually, it’s recommended to incubate for 45-60 mins at room temperature.
Notice: 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.
5.Wash the slides three times with PBST.
6.Proceed to counterstaining, mounting and imaging.
Stage 6 Detection
After incubation with antibodies, you’re now ready to image your slides according to the procedures below.
【Colorimetric】
Materials required
- Tissue slides stained with enzyme-conjugated antibody
- Chromogenic substrate (examples for HRP: DAB D573235, AEC A383182)
- Counterstain (optional, example: Mayer’s Hematoxylin H301933)
- Mounting medium
- Sealant (optional for aqueous mounting media, example nail polish or Limonene) (example M292692)
- Coverslip
- Microscope
Steps
1.Immerse the slide in chromogenic substrate solution. And incubate until the desired color is observed.
Notice: Monitor the staining visually during the incubation.
AP blocking can be included here if required.
2.Wash the slides with cold running water to remove excess stain.
3.Immerse the slide in counterstain solution (optional).
3.1 Incubate according to the manufacturer’s guidance at room temperature or until the desired color is observed.
4.Wash the slides with cold running water to remove the excess stain and to blue the hematoxylin.
5.Dehydrate and clear the tissue before applying organic mounting media (optional).
5.1 Perform the following exchange at room temperature, manually in a Coplin’s jar, or in an automated embedding system.
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6.Add a few drops of mounting medium to the slid: let the slide stand at room temperature for 5 mins.
Notice: Use the minimum volume needed to cover the slides.
7.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.
Notice: Be careful not to introduce any bubbles or disturb the sample.
8.Image the slides using a microscope.
Notice: If not using immediately, store slides at 4°C.
【Florescent】
Materials required
- Tissue slides stained with fluorophore-conjugated antibody
- Fluorescent counterstain (optional, example: DAPI D106471)
- Mounting medium suitable for fluorescent detection (example F598330)
- Sealant (example M292692, A598329)
- Coverslip
- Microscope
- Deionized (DI) water
Steps
1.Immerse the slide in counterstain solution according to the manufacturer’s guidance at room temperature or until the desired color is observed (optional) .
Notice: Some mounting media are fortified with a fluorescent counterstain, which eliminates the need for an additional counterstain step.
2.Wash the slides with cold DI water to remove the excess stain.
3.Add a few drops of mounting medium to the slides. Then let slide stand at room temperature for around 5 mins.
Notice: Use the minimum volume needed to mount the slides.
4.Carefully place a coverslip over the slide using forceps.
- If using an aqueous mounting medium, seal the coverslip with limonene or nail polish.
Notice: Be careful not to introduce any bubbles or disturb the sample.
5.Image the slides using a microscope.
Notice: If not using immediately, store slides in the dark at 4°C.
For more product details, please visit the Aladdin Scientific website.