Immunohistochemistry of Frozen Tissue 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

Materials
Tissue, freshly isolated
PBS, pH 7.4
OCT embedding compound
Liquid nitrogen
Poly-L-lysine-coated slides
Acetone (reagent grade) or 4% (w/v) paraformaldehyde/PBS
0.3% (v/v) H2O2/PBS
Blocking solution: 5% (v/v) goat serum or 1% BSA in PBS
Primary antibody diluted in either PBS, 1% (v/v) goat serum/PBS, or 0.1% (v/v) saponin/PBS
Biotinylated secondary antibody
HRPO-conjugated avidin-biotin complex (ABC): e.g., ABC Elite (for use with human tissue) or ABC Standard (for use with mouse tissue; both from Vector Labs)
AEC substrate solution (Vector Labs)
Hematoxylin counterstain (Mayer’s; Sigma)
Aquamount (Shandon/Lipshaw)
Petri dishes
Aluminum foil, cut into ∼3.5 × 3.5–cm pieces and labeled on the outside using a waterproof marker
Cryostat
Coplin jar or staining dish with slide rack
Pap pen (Research Products International or Shandon/Lipshaw)
Staining chamber
Glass coverslips
Clear nail polish (optional)

Freeze the tissue sample
1. Place the freshly isolated tissue in a petri dish containing PBS and cut into the desired number of ∼0.5-cm3 pieces.
2. Place one tissue piece into the center of a labeled, precut piece of aluminum foil.
3. Apply OCT embedding compound to the fragment in sufficient quantity to completely cover the tissue. Fold the foil to create a secure envelope.
OCT embedding compound is used to prevent dehydration at the edges of the sample and to provide greater surface area for manipulation of the frozen tissue.
4. Drop the foil envelope directly into liquid nitrogen. Incubate the specimen 5 to 10 min. Remove the aluminum envelope with forceps and store at −70° or −135°C (for long-term storage).

Cryosection the tissue
5. To prepare the tissue for cryosectioning, bring the aluminum foil envelope containing the tissue to the cryostat on dry ice or in liquid nitrogen, and open the envelope inside the cryostat.
The cryostat should always be at approximately −20°C and, like a refrigerator, should not be turned off except for cleaning.
6. Cover the cryostat chuck with a thin layer of OCT embedding compound. Place the tissue on the OCT-coated chuck in the appropriate orientation. Allow the tissue/OCT/chuck complex to freeze solid (∼10 min at −20°C) inside the cryostat.
Covering the cryostat chuck is particularly important for such tissues as eye or brain where orientation is critical. The temperature should be between −18° and −20°C; any colder or warmer makes cutting the tissue difficult.
7. Set the cryostat to cut 5- to 8-µm-thick sections.
If the tissue contains experimental cells that are particularly large (e.g., brain), 8- to 10-µm-thick sections should be cut.
8. As the ribbon of tissue comes off the frozen block, place a poly-L-lysine-coated slide under the sections and collect a maximum of three sections clustered together on each slide (Fig. 21.4.2B).
Because of the temperature differential, as the section touches the slide, the section will flatten and stick to the slide.
9. Allow the glass slide with the cryostat sections to air dry, preferably overnight at room temperature.
Dried slides are ready for fixation.

Fix sections
10. Immerse the slides 5 min in a Coplin jar or staining dish containing reagent-grade acetone. Allow the slides to air dry 10 min at room temperature.
Acetone fixation should always be done first, because acetone provides optimal retention of antigenic determinants and thus maximal staining. However, acetone fixation often distorts morphology, depending on the specific tissue used. If tissue morphology is critical, paraformaldehyde fixation can be used. Ten minutes in 4% paraformaldehyde/PBS is an alternative, more gentle fixative that has been shown to preserve some degree of antigenicity, although not as well as acetone does. Paraformaldehyde, however, produces superior morphology.
Coplin jars can be used for fixing and washing up to five slides; a staining dish with a slide rack can be used for up to 20 slides. These containers are useful because they expose the slides to a large volume of solution that completely covers the slides.
11. Using a Pap pen, outline the tissue sections on the glass slide.
Encircling the sections using a Pap pen (water repellant wax) creates a boundary that prevents the reagent from spreading over the entire surface of the slide. When the sections are outlined in this manner, small quantities of reagent can be used with minimal risk of drying.
Some antigens are not stable to storage, so it is recommended that slides be stained as soon as possible after preparation to obtain optimal staining. If slides must be stored, storage at −70°C is usually adequate. However, it may be necessary to test slides over time to determine if and when a particular epitope loses its ability to bind antibody.

Stain the tissue
12. Place the slide in a staining chamber and add PBS to the slide. Incubate 5 min at room temperature.
IMPORTANT NOTE: Take special precautions to prevent the slide from drying during the staining procedure; it is best to change reagents on not more than 3 to 4 slides at a time. When in doubt about drying, cover the slide with more reagent. Also, line the staining chamber with paper towels soaked in sterile water to maintain a moist environment.
Use a squeeze bottle or pipet to add the PBS, but do not apply it directly to the tissue, because the force of the liquid could lift up or destroy the tissue.
Perform all incubation and washing procedures in the staining chamber. If it is necessary to interrupt the protocol, the slides may remain in PBS at any wash step for up to 2 hr without deleterious effects.
13. Remove the PBS by tipping the slide so liquid drains into the staining chamber.
14. Treat the slides with 0.3% H2O2/PBS for 5 min or until bubbling stops.
If bubbling has not stopped after 5 min, reapply H2O2/PBS for another 5 min and, if necessary, repeat until bubbling stops.
This incubation is used to eliminate endogenous peroxidase activity; if the enzyme to be used in amplification is not peroxidase, pretreatment with H2O2 is not necessary. Similarly, pretreatment with H2O2 isn’t needed if the tissue has no endogenous peroxidase activity. This can be determined by performing steps 22 and 23 of this protocol on a test slide (this should take ≤20 min). If there is red precipitate on the slide (substrate without antibody), then peroxide pretreatment is necessary.
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 (see Critical Parameters). 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. 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.

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