ROUTINE MAYER'S HEMATOXYLIN AND EOSIN STAIN

ROUTINE MAYER'S HEMATOXYLIN AND EOSIN STAIN

Manual of Histologic Staining Methods of the Armed Forces Institute of Pathology (Third Edition). American Registry of Pathology ( Luna, Lee G., HT(ASCP) (editor)), McGraw Hill Publishers, New York 1960 (Progressive Stain)

Mayor's hematoxylin is used because it eliminates the necessity for differentiation and bluing of the section. It can be considered a progressive stain which produces a stained section with a clearly defined nuclei while the background is completely colorless. The biggest objection to Mayer's hematoxylin as used in the past, has been that stained slides often fade after I to 3 years. This problem can be eliminated, however, when the slides are washed, after the hematoxylin, in running water for a minimum of 20 minutes.

This method gives consistent results even when more than one person stains sections from the same block. Also, slides may be left in the hematoxylin for hours with-out overstaining. Because of the simplicity of the technique, it is possible to teach others to use it within a shorter time as well as a definite reduction in time performance of the stain itself. See Figure 15 for the steps and timing with use of Mayor's Hematoxylin and Eosin procedure.

FIXATION. Any well fixed tissue.

TECHNIQUE. Paraffin, celloidin, or frozen.

SOLUTIONS: MAYER'S HEMATOXYLIN; EOSIN SOLUTIONS; GRAM'S OR LUGOL'S IODINE.

STAINING PROCEDURE

1. Deparaffinize and hydrate to water.

2. If sections are Zenker-fixed, remove the mercuric chloride crystals with iodine and clear with sodium thiosulphate (hypo) (seepage 41). 3. Mayer's hematoxylin for 15 minutes.

4. Wash in running tap water for 20 minutes.

5. Counterstain with eosin from 15 seconds to 2 minutes depending on the age of the eosin, and the depth of the counterstain desired. For even staining results dip slides several times before allowing them to set in the eosin for the desired time.

6. Dehydrate in 95% and absolute alcohols, two changes of 2 minutes each or until excess eosin is removed. Check under microscope.

7. Clear in xylene, two changes of 2 minutes each.

8. Mount in Permount or Histoclad.

RESULTS

Nuclei - blue - with some metachromasia.

Cytoplasm - various shades of pink-identifying different tissue components.

REMARKS The adhesives used to attach sections onto the slides (gelatin, egg albumen) will sometimes stain, in areas around the section, with Mayor's hematoxylin. This will give the slides a slightly dark appearance but in no way affects the nuclear staining. To remedy this, use 10-12% glacial acetic acid in 95% alcohol, to "clean" the slides after Mayor's hematoxylin. Following with a few dips in saturated aqueous lithium carbonate, the nuclei will blue immediately. This is optional, for the 20-minute wash in running water is sufficient to blue the nuclei. This step will in no way alter or minimize the staining of the nuclei.

REFERENCE. Histopathology Laboratories, Armed Forces Institute of Pathology, Washington, D.C. 20305. Gomori's One-Step Trichrome Stain

Gomori,Sheehan and Hrapchak IN: Histotechnology A Self-Instructional Text ASCP Press. American Society of Clinical Pathologists Chicago 1990

Purpose To identify an increase in collagenous connective tissue fibers, or to differentiate between collagen and smooth muscle fibers.

Principle In the one-step trichrome procedure, a plasma stain (chromotrope 2R) and a connective tissue fiber stain (fast green FCF, light green, or aniline blue) are combined in a solution of phosphotungstic acid to which glacial acetic acid has been added. Phosphotungstic acid favors the red staining of muscle and cytoplasm. The tungstate ion is specifically taken up by collagen, and the connective tissue fiber stain is subsequently bound to this complex, coloring the collagen green or blue, depending on the counterstain used.

Fixative Any well-fixed tissue may be used. Bouin's solution is used as a mordant to intensify the color reactions.

Equipment 56 to 58 °C oven, Coplin jars, Erlenmeyer flasks, graduated cylinders, pipettes.

Technique Cut paraffin sections at 4 to 5 mm.

Quality Control Practically every tissue has an internal control, so no other control sections are needed; however, if a control is desired, uterus, small intestine, appendix, or fallopian tube will provide good material.

Reagents

Bouin's Solution

Picric acid, saturated aqueous solution ….. 75.0 mL

Formaldehyde, 37% to 40% ….. 25.0 mL

Glacial acetic acid ….. 5.0 mL

Weigert's Iron Hematoxylin

Solution A

Hematoxylin..... 10.0 g

Alcohol, 95% ..... 1,000.0 mL

Solution B

Distilled water ..... 475.0 mL

Hydrochloric acid, concentrated ..... 5.0 mL

Ferric chloride, 29% solution ..... 20.0 mL

Working Solution

Mix equal parts of solutions A and B.

Gomori's Trichrome Stain

Chromotrope 2R (Baker No. 5-F703) . . . . . 0.6 g

Fast green FCF, light green or aniline blue ..... 0.3 g

Phosphotungstic acid ..... 0.8 g

Glacial acetic acid . . . . . 1.0 mL

Distilled water ..... 100.0 mL

Store this solution in the refrigerator.

0.5% Acetic Acid Solution

Glacial acetic acid ..... 0.5 mL

Distilled water ..... 99.5 mL

Procedure

1. Deparaffinize sections and hydrate to distilled water. 2. Rinse well in distilled water.

3. Mordant sections in Bouin's solution for I hour at 56 °C.

4. Remove slides from the oven, allow to cool, and wash in running water until the yellow color disappears.

5. Rinse in distilled water.

6. Stain sections in Weigert's hematoxylin for 10 minutes.

7. Wash in running water for 10 minutes.

8. Stain sections for 15 to 20 minutes in Gomori's trichrome stain.

9. Differentiate for 2 minutes in 0.5% acetic acid.

10. Dehydrate, clear, and mount with synthetic resin.

Results

(Plate 8-5, page 267

Nuclei..... black

Cytoplasm, keratin, muscle fibers ..... red

Collagen and mucus ..... green or blue

Notes

1. Sweat et al state that coloration of fine connective tissue fibers is affected by the dye solution pH, with maximum binding occurring around pH 1.3. The pH of Gomori's trichrome is about 2.5, which decreases affinity for anions by approximately 50%, so these investigators suggest that by replacing the acetic acid with hydrochloric acid, a pH of approximately 1.3 can be obtained. The intensity of coloration of the fine connective tissue fibers can be varied by altering the pH.

Verhoeff's Elastic Stain

Mallory, Sheehan and Hrapchak IN Histotechnology A Self-Instructional Text ASCP Press. American Society of Clinical Pathologists, Chicago 1990

Purpose Elastic fiber techniques are used for the demonstration of pathologic changes in elastic fibers. These include atrophy of the elastic tissue, thinning or loss that may result from arteriosclerotic changes, and reduplication, breaks, or splitting that may result from other vascular diseases. The techniques also may be used to demonstrate normal elastic tissue, as in the identification of veins and arteries, and to determine whether or not the blood vessels have been invaded by tumor.

Principle The tissue is overstained with a soluble lake of hematoxylin-ferric chloride-iodine. Both ferric chloride and iodine serve as mordants, but they also have an oxidizing function that assists in converting hematoxylin to hematein. The mechanism of dye binding is probably by formation of hydrogen bonds, but the exact chemical groups reacting with the hematoxylin have not been identified. This method requires that the sections be overstained and then differentiated, so it is regressive. Differentiation is accomplished by using excess mordant, or ferric chloride, to break the tissue-mordant-dye complex. The dye will be attracted to the larger amount of mordant in the differentiating solution and will be removed from the tissue. The elastic tissue has the strongest affinity for the iron-hematoxylin complex and will retain the dye longer than the other tissue elements. This allows other elements to be decolorized and the elastic fibers to remain stained. Sodium thiosulfate is used to remove excess iodine. Van Gieson's solution is the most commonly used counterstain, but others may be used.

Fixative Any well-fixed tissue may be used.

Equipment Mechanical stirrer, Coplin jars, Erlenmeyer flasks, graduated cylinders.

Technique Cut paraffin sections at 4 to 5 mm.

Quality Control Use a section of aorta embedded on edge, or a cross section of a large artery.

Reagents Lugol's Iodine

Iodine..... 10.0 g

Potassium iodide ..... 20.0 g Distilled water ..... 1,000.0 mL

Put the iodine and potassium iodide in a flask with 200 mL of the water. Stir on a mechanical stirrer until the iodine dissolves and then add the remaining water.

100% Ferric Chloride

Ferric chloride ..... 50.0 g

Distilled water ..... 500.0 mL

Store in the refrigerator.

Verhoeff’s Elastic Stain

Prepare fresh each time and mix in order:

Hematoxylin, 5% in 95% alcohol (may be kept as a stock solution) ..... 30.0 mL

Ferric chloride, 10% solution ..... 12.0 mL

Lugol's iodine ..... 12.0 mL

Van Gieson's Solution

Acid fuchsin, 1% aqueous ..... 20.0 mL

Picric acid, saturated solution (14 g/L) ..... 380.0 mL

5% Sodium Thiosulfate

Sodium thiosulfate ..... 50.0 g Distilled water ..... 1,000.0 mL Procedure 1. Deparaffinize sections and hydrate to distilled water.

2. Place sections in Verhoeff’s elastic tissue stain for I hour.

3. Wash in two changes of distilled water.

4. Differentiate sections microscopically in 2% ferric chloride until the elastic fibers are distinct and the background is colorless to light gray. If the sections are differentiated too far, restain. 5. Rinse sections in distilled water.

6. Place sections in sodium thiosulfate for I minute.

7. Wash in running tap water for 5 minutes.

8. Counterstain sections in van Gieson's stain for 1 minute.

9. Differentiate in 95% alcohol.

10. Dehydrate in absolute alcohol, clear in xylene, and mount with synthetic resin.

Results

(Plates 8-7 through 8-9, page 268)

Elastic fibers ..... blue-black to black

Nuclei ..... blue to black

Collagen..... red

Other tissue elements ..... yellow

Notes

1. It is easy to overdifferentiate this stain (Plate 8-8, page 268). If the background is completely colorless, so that a clear yellow counterstain is obtained, the section may be overdifferentiated. It is probably better to err on the side of underdifferentiation.

2. Overdifferentiated sections may be restained at any step provided they have not been treated with alcohol.

3. Do not prolong staining with van Gieson's solution as picric acid also will differentiate the stain further.

4. It is not necessary to remove mercury deposits before staining, as they will be removed by the staining solution.

5. The preparation of van Gieson's solution is critical for proper differentiation of muscle and collagen. If the picric acid is not saturated, collagen will not stain red, and cytoplasm, muscle, and collagen may all stain the same color.

6. To prepare the Verhoeff’s elastic staining solution, the reagents must be added in the order given, with mixing after each addition, or poor staining may result.

7. The staining jar that contained the Verhoeff’s solution may be cleaned easily by transferring the 2% ferric chloride to the jar for a few minutes before discarding the solution.

For optimum results, slides must be individually differentiated, as the time of differentiation is somewhat dependent on the amount of elastic tissue present. Do not depend on the control for timing the differentiation of all sections.

PAS Reaction With Diastase Digestion

Principle This is a very sensitive histochemical method for glycogen. Diastase and a-amylase act on glycogen to depolymerize it into smaller sugar units (maltose and glucose) that are washed out of the section. The Schiff reaction has been described in the PAS procedure.

Fixative 10% neutral buffered formalin, formalin alcohol, or absolute alcohol.

Equipment Hot plate, pH meter, Coplin jars, balance, Erlenmeyer flasks, graduated cylinders, filter paper.

Technique Gut two paraffin sections at 4 to 5 mm. Label one section "with" and one section "without."

Quality Control Two control sections of liver containing glycogen must be used, one labeled "with" and one labeled "without."

Reagents

0.5% Periodic Acid

Periodic acid ..... 2.5 g

Distilled water ..... 500.0 mL

IN Hydrochloric Acid

Hydrochloric acid, concentrated (specific gravity, 1.19) ..... 83.5 mL Distilled water ..... 916.5 mL

Add the acid to the water and mix well.

Schiff Reagent Distilled water ..... 800.0 mL Basic fuchsin ..... 4.0 g

Sodium rnetabisulfite ..... 4.0 g IN hydrochloric acid ..... 80.0 mL

Heat the water to the boiling point. Remove from flame, add the basic fuchsin, and reheat to the boiling point. Cool the solution to 50 °C and Filter. Add 80.0 mL of IN HCI, cool completely, and add 4.0 g of sodium metabisulfite. Place the solution in the dark overnight. The solution should be light amber after standing. Add 2.0 g of activated charcoal and shake for I minute. Filter and store the solution in the refrigerator.

Test for Quality of Schiff Reagent

(See the PAS procedure in this chapter.)

0.55% Potassium Metabisulfite

Potassium metabisulfite ..... 2.75 g

Distilled water ..... 500.0 mL

Malt Diastase Solution

Diastase of malt ..... 0.1 g

Phosphate buffer, pH 6.0 ..... 100.0 mL

Phosphate buffer, pH 6.0

Sodium chloride ..... 8.0 g

Sodium phosphate, monobasic . .. .. 1.97 g

Distilled water . .. .. 100.0 mL

Adjust pH to 6.0 if necessary.

Procedure

1. Deparaffinize and hydrate slides to distilled water.

2. Place the sections labeled "with" in preheated diastase solution at 37 °C for I hour. Hold the sections labeled "without" in distilled water.

3. Wash in running water for 5 minutes.

4. Place all sections ("with" and "without") in 0.5% periodic acid solution for 5 minutes.

5. Wash in three changes of distilled water.

6. Place in Schiff reagent for 15 minutes.

7. Wash for 1 minute in each of two jars of 0.55% potassium metabisulfite to remove excess stain.

8. Wash in running tap water for 10 minutes to develop full color.

9. Counterstain ½ minute in Harris's hematoxylin with acetic acid (2 mL acetic acid/48 mL hematoxylin).

10. Wash well in running water to blue the hematoxylin.

11. Dehydrate with two changes each of 95% and absolute alcohol, clear with xylene, and mount with synthetic resin.

Results

(Plates 7-2 and 7-3, page 264)

Glycogen will stain bright rose red on the section labeled "without" and will be absent from the section labeled "with."

Notes

Malt diastase, containing both a- and (3-amylase, is commonly used for digestion but tends to loosen the sections. For this reason as well as the decreased digestion time, many laboratories prefer to use human saliva, which contains only a-amylase. If preferred, digest with saliva for 20 minutes at room temperature.
Glycogen fixed in picric acid-containing fixatives may be more resistant to diastase digestion than when digestion follows other fixatives (Sheehan and Hrapchak).

Routine Paraffin Cytoplasmic,

Membrane Antigens and Nuclear Antigens

PurposeImmunohistochemistry is used to identify a long list of antigens. It is a very effective and flexible technique that is dependent upon the specificity and sensitivity of the primary antibody. If one has an antibody that identifies the fixed, processed antigen, any number of secondary antibodies and/or amplification techniques can be used to identify the antigen in paraffin sections.

Principle The tissue is processed and unstained paraffin sections are mounted on glass slides. Most procedures call for "antigen retrieval" using microwave ovens and citrate buffers. The incubation in methanol and hydrogen peroxide is designed to "kill" the endogenous peroxidases and eliminate background staining. The appropriate dilution of primary antibody is placed in solution on the deparaffinized sections and exposed of an appropriate time interval (depending on the affinity and avidity of the antibody and the preservation and concentration of the antigen). The secondary antibody systems are then placed on the slide. The procedures provided below illustrate the use of the ABC kits but many different protocols, signal amplification systems and indicator systems are available. The new mouse-on-mouse systems permit the use of some mouse monoclonal antibodies on mouse tissue.

Fixative In general, tissue fixed in paraformaldehyde for less than 48 hours provides the best results. However, different antigens will require different protocols. Post fixation of fomalin or paraformaldehyde fixed tissue with B5 or other heavy metal fixative can "retrieve" some antigens.

The protocol for Mouse-on-Mouse used on the slide set is found by clicking here.

Paraffin mounted slides

1. Xylene 5 min.

2. Xylene 5 min.

3. Xylene/Lugol’s 4 min. *

4. Absolute ETOH 3 min.

5. Absolute ETOH 3 min.

6. MEOH/H2O2(0.3%) 20 min.

7. Absolute ETOH 1 min.

8. 95% ETOH 2 min.

9. 70% ETOH 2 min.

10. H2O (running) 2 min.

11. Dist. H2O 2 min.

12. PBS (x2) 5 min.

13. Normal Equine Serum 10% 20 min.

14. Primary abs. (Dilutions vary) Overnight

15. PBS (x2) 5 min.

Equine Anti-Mouse Biotin Conjugate 60 min.

17. PBS (x2) 5 min.

18. ABC - Elite 30 min.

19. PBS (x2) 5 min.

20. DAB (Vector-Brown) 3-5 min.

22. Mayer’s Hematoxylin 1 min.

23. Tap H2O 5-10 min.

24. Dehydrate, clear and coverslip

*B5 Fixed Tissue only
NOTE: Tissue sections are placed in a 55 Celsius oven for 30 minutes or overnight.

Immunoperoxidase Staining (Nuclear Antigens)

1. Xylene 5 min. RoomTemp.

2. Xylene 5 min. RoomTemp.

3. ETOH 3 min. RoomTemp.

4. ETOH 3 min. RoomTemp.

5. H202 + 3% Methanol 20 min. RoomTemp..

6. ETOH 2 min. RoomTemp.

7. ETOH 2 min. RoomTemp..

8. H2O tap 5 min. RoomTemp..

9. distilled H2O 5 min. RoomTemp..

10. Citrate Buffer 4 min. microwave*

11. Citrate Buffer 4 min. microwave*

12. Citrate Buffer 4 min. microwave

13. Cool down 15 min. RoomTemp.

14. PBS 5 min. RoomTemp..

15. PBS 5 min. RoomTemp.

16. N horse serum 20 min. RoomTemp.

17. Primary antibody overnight 4 C

18. PBS 5 min. RoomTemp.

19. PBS 5 min. RoomTemp.

20. Secondary (BHAM) 1:800 60 min. RoomTemp.

21. PBS 5 min. RoomTemp.

22. PBS 5 min. RoomTemp.

23. Tertiary ABC 1:50 30 min. RoomTemp.

24. PBS 5 min. RoomTemp.

25. PBS 5 min. RoomTemp.

26. DAB 3-5 min. RoomTemp.

27. H2O running tap 5 min. RoomTemp.

28. Hematoxylin-Mayers 30 seconds RoomTemp.

29. H2O running tap

30. Dehydrate, clear, coverslip