IHC / Immunohistochemistry Protocol

Chromogenic Immunohistochemistry Protocol for Paraffin Embedded Tissues.

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Introduction

Immunohistochemistry (IHC) is a common approach for visualizing localization of specific protein expression within tissue sections using antibodies.  The following immunohistochemistry protocol will discuss immunohistochemistry in formalin-fixed, paraffin-embedded tissues.  In IHC, tissue samples are chemically fixed to preserve signaling interactions.  These samples are then embedded in paraffin wax or frozen, thinly sectioned, and mounted to a slide for antibody labeling and analysis.  In this immunohistochemistry protocol, these samples are then exposed to a chromogenic substrate such as DAB which causes a color change that can be observed under a microscope.

Materials

1.       Paraffin-Fixed Tissue Samples

2.       Paraffin-Wax (eg. Paraplast® from Leica)

3.       Charged Slides (eg. Apex BOND from Leica)

4.       Xylene

5.       Histology Grade Ethanol (Anhydrous, Denatured 100% and 95%)

6.       Wash Buffer (Tris-Buffered Saline with Tween® 20 (TBST))

7.       Primary Antibody Diluent (eg. 1% BSA (block and stabilize), 0.1% Fish Gelatin (block), and 0.5% Triton-X (penetrant), in PBS)

8.       Secondary Antibody Diluent (eg. 0.5% Triton-X (penetrant) in PBS)

9.       A counterstain such as hematoxylin

10.   Antigen Retrieval Solutions (Generally 10 mM sodium citrate buffer pH 6.0 is preferred)

11.   3% Hydrogen Peroxide (H2O2)

12.   Blocking Solution (TBST with 5% Normal Serum from same host as antibody)

13.   Primary Antibody

14.   HRP-conjugated secondary antibody (Specific to host of primary antibody, eg. Anti-mouse or anti-rabbit).

         15.   Substrate (eg. 3,3-diaminobenzadine (DAB))

Methods

Tissue Preparation

1.       Place the paraffin-fixed tissue sample in a mold with a residual volume of liquid paraffin.  Cool until tissue is immobilized and then fill with liquid paraffin.  Cool.

2.       Section tissues (generally 4-6 µm) on a microtome and float sections in a water bath.

 3.        Mount the sections to the charged slides and allow to dry overnight.  The charged slides ensure well adequately tissue sections with well preserved morphology.

Paraffin Removal and Rehydration

1.       Remove paraffin by immersing the tissue in 3 separate containers of xylene for 5 minutes each.  This step is critical as incomplete paraffin removal results in inconsistent staining.

 2.       To rehydrate tissues, rinse twice in 100% ethanol and twice in 95% ethanol (ensure fresh ethanol is used for each step) for 10 minutes each.  Then rinse twice in dH2O (5 minutes each rinse).

Antigen Retrieval

Note:  Cross-linking created during formalin fixation must be reversed in order to expose antigens to which antibodies bind.

1.       Using a microwave, bring tissue sections to a boil for 10 minutes in 10 mM sodium citrate buffer.  Care must be taken to avoid a vigorous boil.  Optimize conditions (power settings) on microwave to ensure a low boil is gradually reached using blank slides prior to treating samples.  Allow 20 minutes for samples to cool to room temperature before proceeding.  Other immunohistochemistry protocols may recommend different buffers or heating methods.  Refer to our datasheets for specific immunohistochemistry protocol modifications required for particular targets.

Chromogenic Staining

1.       Wash sections twice in dH2O (5 minutes each rinse).

2.       Incubate in 3% H2O2 to quench any endogenous peroxidase activity which may contribute to background and decrease signal:noise.

3.       Wash twice in dH2O (5 minutes each rinse) and again in wash buffer (5 minutes).

4.       Incubate for 1 hr. with ~200 µl of blocking solution at room temperature to reduce non-specific binding of the antibodies.

5.       While blocking, dilute antibody to 2-4 µg/ml in primary antibody diluent (enough for 100-400 µl of dilute antibody per sample with excess for loss attributable to pipetting).  We recommend piloting several dilutions prior to running a full experiment to determine the optimal dilution for your samples.

6.       Add 100-400 µl of dilute antibody to each sample and incubate overnight in a refrigerator at 4°C.

7.       Equilibrate the secondary antibody to room temperature.

8.       Dilute secondary antibody to 0.5-5 µg/ml in secondary antibody diluent (refer to manufacturer datasheet for recommended dilution).

9.       Wash three times with wash buffer (5 minutes each rinse).

10.   Add 100-400 ul of dilute secondary antibody to each sample and incubate 1 hr. at room temperature.

11.   Wash three times with wash buffer (5 minutes each rinse).

12.   Add 100-400 ul of DAB substrate to each sample and incubate 2-10 minutes monitoring closely.

13.   Immerse slides for 5 minutes in dH2O to stop the reaction.

14.   If desired, apply counterstain such as hematoxylin to contrast tissue morphology.

15.   Wash sections twice in dH2O (5 minutes each rinse).

Dehydrating Samples and Mounting Sections

1.     This is essentially a fast reversal of the rehydration step above.  Place each slide in 95% ethanol twice for ten seconds each (ensure fresh ethanol is used for each step).

2.  Place each slide in a container of 100% ethanol twice for ten seconds each.

 3.   Place each slide in a container of xylene twice for 10 seconds each.

 4.   Mount cover-slips using a mounting medium such as clear nail polish.