Immunohistochemistry of Fixed Paraffin-Embedded Sections

Fixation, Embedding, and Sectioning of Tissues, Embryos, and Single Cells
Florence Hofman

University of Southern California, Los Angeles, California

John E. Coligan, Ada M. Kruisbeek, David H. Margulies, Ethan M. Shevach, and Warren Strober (eds.)
Current Protocols in Immunology
Copyright © 2003 John Wiley & Sons, Inc. All rights reserved.
DOI: 10.1002/0471142735.im2104s49
Online Posting Date: August, 2002
Print Publication Date: June, 2002

Additional Materials
Tissue fragment or paraffin-embedded section
10% (v/v) neutral buffered formalin or 4% (w/v) paraformaldehyde in PBS, pH 7.4
100%, 95%, and 80% ethanol
0.3% (v/v) H2O2/methanol

1. Immerse the tissue fragment in 10% neutral buffered formalin or 4% paraformaldehyde in PBS for 6 to 8 hrs, depending on the size of tissue.
The optimal size for the tissue fragment is 5 × 5 × 3 mm.
If tissue is already embedded in paraffin, begin with step 4.
2. Process the tissue: Dehydrate 15 min in 80% ethanol; 15 and then 20 min in 95% ethanol; three times, 20 min each, in 100% ethanol; and finally, three times, 20 min each, in xylenes. Embed in paraffin, section 5- to 8-µm thick, and place the sections on poly-L-lysine-coated slides.
Tissue processing and slide preparation are standard and usually performed in a dedicated histology laboratory.
3. After the tissue sections are on the slide, dry slides 1 hr in a 60° oven.
This drying procedure removes water from beneath the tissue section and makes the section adhere better to the slide.
4. Deparaffinize the slide by placing it for two 5-min changes each in Histoclear, 100% ethanol, and 95% ethanol. Between changes, quickly blot the edge of the slide on a paper towel to remove excess liquid, but do not allow the slide to dry.
Histoclear is a substitute for potentially toxic xylenes.
Immersion steps are performed using either Coplin jars or staining dishes with slide racks, depending on the number of slides that are being processed. The containers are filled so that the slides are completely immersed when they are inside the container. The solutions are reusable; however, they must be replaced once or twice a week depending on the number of slides processed and the amount of liquid carried over into the containers with the slides. Usually the solutions are changed after every 50 slides. However, because these reagents absorb water from the atmosphere, they should be changed at least every 2 weeks no matter how many slides have been treated.
5. To block endogenous peroxidase, incubate the slide 10 min in 0.3% H2O2/methanol.
This step should always be performed when peroxidase is the enzyme used for amplification; it should be omitted when other enzymes are used for detection.
6. Wash slides twice in PBS, 10 min each at room temperature.

15. Add blocking solution to the slide and incubate 15 min. Tilt the slide to remove the solution. Before going to the next step (applying the antibody), remove as much of the liquid as possible, without touching the tissue or letting the section dry.
Remove excess liquid by blotting the edge of the slide on a paper towel so the antibody will not be unnecessarily diluted when it is added to the slide.
16. Apply diluted primary antibody in sufficient quantity to cover the tissue within the area marked by the wax pen. Incubate 30 to 60 min at room temperature.
Depending on the specificity of the primary antibody, it may be necessary to optimize working dilution.Optimization may be performed using cell suspensions.
To detect cytoplasmic antigens, the primary antibody may have to be diluted in 0.1% Triton X-100 or saponin (see Critical Parameters). Detergent should only be used to provide cytoplasmic access if a positive control tissue exposed to primary antibody diluted in PBS gives a negative result. If detergent is added to the antibody, it should also be added to the PBS used for washing the slides after antibody incubations.
Commercial ABC kits are sold with species-specific secondary antibodies. The kits include specific guidelines for appropriate working dilutions.
17. Wash the slide twice in PBS, 10 min each.
18. Apply diluted biotinylated secondary antibody to the tissue within the wax outline. Incubate 30 min at room temperature.
The secondary antibody should recognize immunoglobulin (Ig) from the species that is the source of primary antibody (e.g., biotinylated goat anti-mouse is used to recognize a mouse monoclonal primary antibody). Secondary rodent-specific polyclonal antisera may need to be preabsorbed against normal rodent serum to remove background reactivity with rodent Ig antigens.
19. Wash the slide twice in PBS, 10 min each.

Detect bound antibodies
20. Incubate the slide 20 min in HRPO-conjugated avidin-biotin complex (ABC).
For human tissue the ABC Elite kit is ideal because of its superior sensitivity. For mouse tissue the ABC Standard kit is preferred, because enhanced sensitivity may bring out unwanted background.
21. Wash the slide twice in PBS, 10 min each.
22. Add AEC substrate solution to the slide and incubate 5 to 10 min. Check the slide after 5 min for color development.
Always prepare the working AEC solution immediately before adding it to the slide, and add H2O2 as the final ingredient. If this solution is not used within 5 min of preparation, staining intensity will be significantly diminished.
To examine the slide, place a white paper towel underneath it and look for red color. If the slide looks bright red, stop the reaction immediately by washing the slide in tap water. To examine the slide more closely, tilt it to remove most of the liquid, then add a coverslip and examine the slide using a light microscope. If the positive control slide is understained, reapply a freshly prepared AEC substrate solution.
23. Transfer the slide to a Coplin jar or staining dish and wash 10 min in running tap water. Direct the stream of water at a corner of the container to provide good circulation and to avoid contact with the tissue on the slide.

Counterstain tissue
24. Immerse slide in hematoxylin counterstain for 30 sec to 2 min.
Hematoxylin is reusable and should be kept in a covered staining dish on the bench top. However, carryover of water from the slides into the hematoxylin will eventually dilute the stain. Thus, after five to ten uses the incubation time required will steadily increase. When nuclear staining is no longer crisp and clear, the stain should be replaced with fresh hematoxylin.
25. Wash the slide 10 min in tap water.
26. Place 1 to 2 drops of Aquamount on the tissue.
27. Place a coverslip over the tissue and, if desired, seal with clear nail polish to prevent drying. Allow the nail polish to dry thoroughly before examining the slide under a light microscope.
Sealed slides can be stored indefinitely.

This general protocol is is intended for use as a reference as a courtesy to our customers. Optimal concentrations, experimental conditions and experimental processes are to be determined by the individual user. No guarantee of performance using the above procedure is expressed or implied. Molecular Innovations does not currently perform immunohistochemistry in-house and can not provide further technical support for this application.

Did You Find This Article Helpful?

Yes - 3 visitors found this post helpful
No - 0 visitors found this post was not helpful