IHC Protocol for Paraffin Embedded Sections

IHC Technical Resources

Protocols, optimization tips,
troubleshooting guides,
and more for IHC.

Troubleshooting guides

Troubleshooting guides

Download troubleshooting
handbooks for IHC, Western
blot and ELISA for FREE.

Summary workflow chart for IHC protocol for Paraffin embedded Sections:

IHC Workflow (Paraffin Sections) with Applicable Boster’s Reagents

  1. Tissue Preparation

    1. Paraformaldehyde Cooling and Dehydration

      • Harvest fresh tissue and place it in a dish filled with ice-cold PBS buffer
      • Wash the tissue thoroughly with PBS to remove blood (Use forceps to remove connective tissues)
      • Cut the tissue into slices of thickness of 3 mm or less
      • Immerse the slices in 4% paraformaldehyde at room temperature for 8 min
      • Immerse the slices in 4% paraformaldehyde (pre-cool at 4°C) for 6 to 7 hrs. The paraformaldehyde volume should be 20X greater than the tissue volume by weight
      • Wash the tissue 3X with PBS (1 min each)
      • Dehydrate the tissue by immersing the tissue sequentially as follows:
        • 1X into 80% ethanol (1 hr at 4°C)
        • 1X into 90% ethanol (1 hr at 4°C)
        • 3X into 95% ethanol (1 hr each at 4°C)
        • 3X into 100% ethanol (1 hr each at 4°C)
        • 3X into dimethylbenzene (0.5 hr each at room temperature)
    2. Liquid Paraffin Section

      • Prepare the first portion of liquid paraffin in a suitable bath and allow the paraffin to reach and maintain at 60°C
      • Immerse the tissue 2X into the paraffin bath (2 hrs each)
      • Prepare the second portion of liquid paraffin in a suitable bath and allow the paraffin to reach and maintain at 60°C
      • Pour the second portion of paraffin into a mold
      • Quickly transport the tissue from the paraffin bath to the mold with paraffin
      • Incubate the tissue at room temperature until it coagulates
      • Store the tissue at 4°C
    3. Section Slicing and Incubation

      • Secure the paraffin section on slicer
      • Slice one to two pieces of section to adjust the slicer so that the section and blade are parallel
      • Slice the remaining section carefully with ~5 µm thick
      • Incubate the sliced section in 40 to 50°C water to unfold
      • Mount the tissue section onto Poly-Lysine or APES coated glass slides
      • Incubate the slides overnight at 37°C

    Note: This fixation procedure using paraformaldehyde and formalin fixatives may cause autofluorescence in the green spectrum. In this case, you may try fluorophores in the (i) red range or (ii) infrared range if you have an infrared detection system.

  2. Dewaxing/Deparaffinization

    • Prepare the following reagents:
      • 90% dimethylbenzene
      • 95% dimethylbenzene
      • 100% dimethylbenzene
      • 90% ethanol
      • 95% ethanol
      • 100% ethanol
    • Sequentially immerse paraffin sections into:
      • 90% dimethylbenzene (for 7 min)
      • 95% dimethylbenzene (for 7 min)
      • 100% dimethylbenzene (for 7 min)
      • 90% ethanol (for 7 min)
      • 95% ethanol (for 7 min)
      • 100% ethanol (for 7 min)
    • Wash the slides with water to remove ethanol

    Note: The process of dewaxing should be done in a fume hood at room temperature in summer. When the temperature is lower than 18°C, it is recommended to dewax at 50°C.

  3. Inactivation

    • Immerse dewaxed paraffin section into the 3% H2O2 at room temperature for 10 min
    • Wash the section 3X to 5X with distilled water (total 3 to 5 min)
  4. Antigen Retrieval (Heat Induced Epitope Retrieval: HIER)

    • Immerse the paraffin sections in citrate buffer
    • Heat the buffer in microwave and turn it off when the buffer has boiled
    • Keep the boiled buffer in microwave for 5 to 10 min
    • Repeat the heating as outlined above 1X to 2X
    • Cool the slides until it reaches room temperature
    • Wash the sections 1X to 2X with PBS
  5. Blocking

    • Add 5% BSA blocking solution or normal goat serum to the HIER treated samples
    • Incubate the samples at 37°C for 30 min
    • Discard extra liquid (No washing required)
  6. Primary Antibody Incubation

    • Dilute primary antibody with antibody diluent to the concentration recommended by the antibody manufacturer
    • Add the diluted antibody to the samples and incubate overnight at 4℃ or at 37℃ for 1 hour
    • Wash the samples 2X with PBS (20 min each)
  7. Secondary Antibody Incubation

    • Dilute biotinylated secondary antibody with antibody diluent to the concentration recommended by the antibody manufacturer
    • Add the diluted antibody to the samples and incubate at 37°C for 30 min
    • Wash the samples 2X with PBS (20 min each)
  8. Staining

    • Add Strept-Avidin Biotin Complex (SABC) HRP- or AP-conjugated reagents to the samples
    • Incubate the samples at 37°C for 30 min
    • Wash the samples 3X with PBS (20 min each)
    • Add a suitable amount of DAB reagent to the samples and incubate in dark at room temperature for 10 to 30 min
    • Monitor the tissue staining intensity under a bright-field microscope*
    • Wash the samples 3X to 5X with distilled water
    • Counterstain (if necessary)
      • Add haematoxylin to the sample
      • Dehydrate
      • Immerse the paraffin sections 2X in dimethylbenzene (7 min each)
    • Check the tissue staining intensity under a bright-field microscope

    *If the staining background is too high, wash the section 4X with 0.01-0.02% TWEEN 20 PBS and 2X with pure PBS after the SABC reaction and before DAB staining. Then use DAB to stain the samples.