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Immunohistochemistry Paraffin

IHC-P

Immunohistochemistry (IHC) is an assay typically used to detect antigens, via labelled antibodies, in biological samples. It can be used to analyse the distribution, abundance and localisation of proteins, glycans and other small or non-biological molecules; such results offer insight into cellular structure and mechanisms. There are two detection methods for immunohistochemistry: colorimetric and fluorescent. Fluorescent detection uses fluoresence-labelled antibodies, and the method is more commonly referred to as Immunofluorescence (IF). Colorimetric detection uses enzyme-conjugated antibodies that produce a coloured precipitate.

See below for a general protocol for the immunohistochemistry staining method SABC (Strept Avidin Biotin-peroxidase Complex), for paraffin sections. Streptavidin labels the target protein through the combination of its carboxyl and the amidogen of the protein. It has high sensitivity and low non-specific binding to tissues and cells, resulting in very low background.

A. Preparation

Sample Preparation

  1. Wash fresh tissue with PBS to remove blood. Remove connective tissue using tweezers, and carve tissue into small pieces. Immerse into 4% Paraformaldehyde for 8-10 mins.
  2. Fix tissue by immersion in pre-cooled 4% Paraformaldehyde (4oC) (20 times the volume of the tissue sample) for 6-7 hrs.
  3. Note: Over-fixation can mask the epitope. Under-fixation can lead to edge staining. 4% Paraformaldehyde can be substituted by Neutral Buffered Formalin (10%) and left for 24-48 hrs.
  4. Wash with PBS for 1 min, 3 times.
  5. Dehydrate tissue by immersion into ethanol (80% ethanol for 1 hr, then 90% ethanol for 1 hr, then 95% ethanol for 1 hr, 3 times, and finally 100% ethanol for 1 hr, 3 times). Carry out dehydration at 4oC.
  6. Clear the tissue by immersion in dimethylbenzene at room temperature for 30 mins, 3 times.
  7. Immerse the tissue into prepared liquid paraffin at 58 to 60oC for 2 hrs, 2 times.
  8. To embed in paraffin block: Pour liquid paraffin, pre-heated to 60oC, into mold. Place tissue into the block as soon as possible, and incubate at room temperature until the paraffin sets. Store at 4oC until sectioning.
  9. To slice paraffin sections: Fix paraffin section on slicer, slice extra paraffin and adjust slicer to ensure paraffin and blade are parallel. Slice paraffin section carefully. Cut 4-6 mm slices.
  10. Incubate sliced paraffin section in a water bath containing distilled water at 40 to 50oC to unfold. Prepare glass slide by immersion into APTES or Poly-Lysine. Dip out paraffin section with prepared glass slide. Incubate slide at 37oC
  11. Dry the paraffin section by incubating for 2 hrs at 60oC.

Dewax

  1. Prepare 3 bottles of 90%, 95% and 100% xylene and 3 bottles of 90%, 95% and 100% ethanol.
  2. Immerse paraffin sections into 3 bottles of xylene orderly (concentration from low to high) for 7 min each. Then immerse into ethanol orderly at room temperature (same order as xylene) for 7 min each.
  3. Wash with water to remove ethanol.
  4. Note: dewaxing should be done in a fume hood at room temperature. If the temperature is below 18 ℃ it is recommended to dewax at 50 ℃.

Inactivation

  1. Prepare a 1:10 solution of 30% H2O2 in distilled water (mix one part 30% H2O2 with nine parts distilled water).
  2. Immerse dewaxed paraffin section into solution at room temperature for 10 min.
  3. Wash with distilled water for several min.

Antigen Retrieval (optional)

  1. Heat repair: Immerse the paraffin sections into citrate buffer (pH 6.0), microwave until boiling then cut off the power, leave in microwave for 10 min. Repeat twice. Cool at room temperature and wash twice by immersing in distilled water.

B. Protocol

  1. Add 5% BSA blocking solution or normal goat serum and incubate at 37 ℃ for 30 mins with shaking.
  2. Dilute primary antibody in blocking buffer. Incubate at 37oC
  3. Wash with PBS for 20 min, 2 times. Add labelled (enzyme- or fluorophore conjugated) secondary antibody and incubate at room temperature in the dark for 1-2 hrs.
  4. Note: When using any primary or fluorochrome-conjugated secondary antibody for the first time, titrate the antibody to determine which dilution allows for the strongest specific signal with the least background for your sample.
  5. Wash with PBS for 20 min, 2 times. Add SABC reagents and incubate at 37 ℃ for 30 mins. Wash with PBS for 20 mins, 3 times.
  6. Note: Prepare solution by mixing 1 ml distilled water with one drop of Reagent A, B, C.
  7. Incubate at room temperature as per instructions.

Counter-staining (if required)

  1. In preparation for ‘counterstaining; with IB4/DAPI or GFAP/DAPI, incubate the sections in PBS with Ca2+, Mg2+, and Mn2+ and 0.1% Triton X-100 for 15 minutes. Rinse the slides through a quick change of PBS.
  2. Incubate slides in 1:10,000 DAPI for 30 min. Wash slides in PBS for 2 mins, 3 times.
  3. Prepare antifade mounting media according to directions. Allow the slides to dry approximately ¾ to completion. Place the coverslip taking care to remove any bubbles formed beneath the coverslips.