6.1 PERIODIC ACID SCHIFF (PAS )FOR THE DEMONSTRATION OF GLYCOGEN
(McManus, 1946) Glycogen is a simple intracytoplasmic polysaccharide found in greatest amount in liver, cardiac and skeletal muscle, with significant quantities also present in hair follicles, endometrial glands, vaginal and cervical epithelium, umbilical cord, neutrophil leucocytes and megakaryocytes.

PRINCIPLE
Tissue containing vicinal glycol groups or their amino or alkylamino derivatives are oxidized by periodic acid to form dialdehydes, which combine with Schiff's reagent to form an insoluble magenta compound. The chromphoric groups of basic fuchsin in Schiff's reagent are broken by sulphuration to form a colourless solution. In the presence of free aldehyde groups an insoluble coloured compound similar to the original dye is formed.

REFERENCES
p 16 Cook H C, Manual of Histological Demonstration Techniques p 188 Bancroft JD & Stevens A, Theory and Practice of Histological Techniques, 2nd ed

SPECIMEN
Standard paraffin section fixed in 10% neutral buffered formalin. CONTROL Normal liver

REAGENTS
(A) 1 % AQUEOUS PERIODIC ACID SCHIFF REAGENT
Commercial Schiff reagent Australian Biostain Product AB1200.0600 (kept in fridge) If unavailable, use the following: Basic fuchsin (CI 42500) Sigma cat # P1528 2.0 g Distilled water 400 mL Potassium or sodium metabisulphite 4.0 g Conc. hydrochloric acid 4 mL Decolourising activated charcoal 0.4 g
1. Bring water to boil, remove flame and add basic fuchsin slowly and stir until dissolved. Warning: - Use a conical flask and always remove from heat after boiling, then add basic fuchsin slowly. If added too fast solution will bubble explosively out of the flask.
2. Cool to 50øC, add metabisulphite. Dissolve and cool to room temperature.
3. Add conc. HCl and mix.
4. Leave overnight in the dark at room temperature. Add charcoal and shake for 1 - 2 minutes. Filter and store at 4øC in dark container. Bring solution to room temperature before use. With use it will eventually be found that the originally colourless to pale yellow solution will turn pink due to loss of sulphur dioxide causing restoration of the basic fuchsin colour. When this happens, the solution should be discarded. If a white precipitate forms it should be filtered out.
NOTES
Schiff Reagent - test of activity: The Schiff reagent may be tested by adding a few drops to 3 to 5 mL of 40 formaldehyde on a petri dish: active Schiff reagent will lead to the rapid development of a reddish purple colour; delayed development of a deep bluish purple indicates that the reagent is going off.

PROCEDURE
1. Deparaffinize and hydrate to distilled water.
2. Oxidise with periodic acid for 5 minutes.
3. Rinse in distilled water.
4. Treat with Schiff's reagent for 5 minutes.
5. Wash in running water for 10 minutes; this intensifies the colour reaction.
6. Stain the nuclei with Mayer's haematoxylin for 1 minute (no need to blue).
7. Wash in water.
8. Dehydrate, clear and mount.

RESULTS
Nuclei Blue
PAS positive material Magenta
Glycogen Cellulose Chitin Neutral mucin Sialomucin : enzyme-labile Starch Sulphated sialomucins Amoebae Fungi Basement membranes
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6.2 PERIODIC ACID SCHIFF (PAS) WITH DIASTASE FOR THE ENZYME EXTRACTION OF GLYCOGEN

PRINCIPLE
Glycogen is digested with certain forms of amylase. Commercially available diastase, which contains both ? and á amylase or salivary amylase from saliva can be used to digest glycogen in tissue sections.

REFERENCES
p 216 Cook H C, Manual of Histological Demonstration Techniques p 188 Bancroft JD & Stevens A, Theory and Practice of Histological Techniques 2nd ed

SPECIMEN
Standard paraffin section fixed in 10% neutral buffered formalin.

CONTROL
Tissue containing glycogen e.g. liver Use two positive controls and two test sections Treat one test section and one positive control with diastase; then stain all sections with PAS.

REAGENTS
(A) AMYLASE SOLUTION
Amylase Type VI-B from porcine pancreas (Sigma A 3176) 0.25g (stored in fridge) Distilled water 10mL If larger quantity required, use 1g Amylase in 40mL distilled water. Prepare fresh just before use. Warm water for 5 sec in microwave oven then add amylase.
NOTE:
AQIS requirements for disposal. Place waste amylase into container and place into yellow biohazard bag for disposal by incineration..
(B) SEE METHOD 6.1 - PAS TECHNIQUE for PAS reagents

PROCEDURE
1. Deparaffinise two positive controls and two test sections and hydrate to distilled water.
2. Treat one control and one test section with Amylase for 25 minutes to digest Glycogen.
3. Wash in running water for 20 minutes.
4. Proceed with P.A.S. technique as follows.
5. Oxidise with periodic acid for 5 minutes.
6. Rinse in distilled water.
7. Treat with Schiff's reagent for 5 minutes.
8. Wash in running water for 10 minutes, this intensifies the colour reaction.
9. Stain the nuclei with Mayer's haematoxylin 1 minute (no need to blue).
10. Wash in water.
11. Dehydrate, clear and mount.

RESULTS
Presence of glycogen will be evidenced by loss of staining after enzyme treatment when compared to the untreated sections.

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6.3 ALCIAN BLUE TECHNIQUE FOR ACID MUCINS
Acid mucin - strongly sulphated connective tissue mucins react at low pH values with suitable cationic dyes and are usually PAS-negative.

PRINCIPLE
Alcian blue is a copper phthalcyanin dye and contains positively charged groups capable of salt linkage with certain polyanions. These polyanions consist of the sulphate and carbocyl radicals of the acid mucins - the phosphate radicals of the nucleic acids do not react. Consequently, only the acid mucins are stained. By varying the pH of the solution of Alcian blue more information can be gained concerning the types of acid mucin present.

REFERENCES
p 200 Cook H C, Manual of Histological Demonstration Techniques p 194 Bancroft JD & Stevens A, Theory and Practice of Histological Techniques, 2nd ed

SPECIMEN
Standard paraffin section fixed in neutral buffered formalin.

CONTROL
Large bowel

REAGENTS
(A) ALCIAN BLUE SOLUTIONS (CI 74240) pH 3.1 - 1% Alcian blue in 0.5% acetic acid Routine: pH 2.5 - 1% Alcian blue in 3.0% acetic acid Use pH 2.5 for routine staining of acid mucins. Solutions keep for 6 months and should be filtered before use. pH 1.0 - 1% Alcian blue in N/10 hydrochloric acid. pH 0.2 - 1% Alcian blue in 10% sulphuric acid.
(B) 3 % AQUEOUS ACETIC ACID (C) 0.5% NEUTRAL RED

PROCEDURE
1. Deparaffinize and hydrate to distilled water.
2. Treat in 3% acetic acid for 2 minutes.
3. Drain off.
4. Stain in 1% alcian blue (pH 2.5) for 30 mins.
5. Wash in water.
6. Counterstain with neutral red lightly (3 mins).
7. Dehydrate rapidly in alcohol.
8. Clear and mount.
 

RESULTS
Alcian blue solution at: pH 3.1 and 2.5 most acid mucins (except some of the strongly sulphated group) Blue
pH 1.0 only weakly and strongly sulphated acid mucins Blue
pH 0.2 only the strongly sulphated acid mucins Blue
Nuclei Red
Background Very pale red or colourless

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6.4 COMBINED ALCIAN BLUE PAS TECHNIQUE FOR ACID AND NEUTRAL MUCINS
(Mowry 1956)

PRINCIPLE
This stain demonstrates the presence of mucins and clearly distinguishes between acid and neutral mucins. By first staining all acid mucins with alcian blue, those acid mucins which are also PAS positive will not react with subsequent PAS reaction, only the neutral mucins.
 

REFERENCES
p 202 Cook H C, Manual of Histological Demonstration Techniques p 197 Bancroft JD & Stevens A, Theory and Practice of Histological Techniques,- 2nd ed SPECIMEN Standard paraffin section fixed in 10% neutral buffered formalin.

CONTROL
Large bowel.

REAGENTS
(A) ALCIAN BLUE SOLUTIONS (CI 74240) 1 g 3% Acetic Acid 100mL Store in fridge.
(B) SCHIFFS REAGENT See P.A.S. technique [ 6.1 ]
(C) 1% PERIODIC ACID
(D) 3% ACETIC ACID

PROCEDURE
1. Deparaffinize and hydrate to distilled water.
2. Acetic acid for 2 mins.
3. Drain.
4. Alcian Blue Solution 10 minutes.
5. Wash in water, then distilled water.
6. Treat with 1% periodic acid for 5 mins.
7. Rinse in distilled water.
8. Treat with Schiffs reagent 5 mins.
9. Wash in running tap water 5 - 10 minutes.
10. Stain nuclei with Mayer's haematoxylin for 1 minute 30 seconds. Nuclear overstaining causes masking of the Alcian Blue staining.
11. Wash in water.
12. Dehydrate, clear and mount.

RESULTS
Acid mucins Blue
Neutral mucins Magenta
Nuclei Pale blue

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6.5 COMBINED ALCIAN BLUE/DIASTASE/PAS FOR ACID & NEUTRAL MUCINS

PRINCIPLE
This stain is useful for the demonstration of acid and neutral mucins. Alcian blue stains acid mucins, subsequent treatment with diastase digests glycogen from the tissue and neutral mucins can then be stained with PAS.

REFERENCES
p 202 & 216 Cook H C, Manual of Histological Demonstration Techniques p 188 & 197 Bancroft JD & Stevens A, Theory and Practice of Histological Techniques, 2nd ed.

SPECIMEN
Standard paraffin section fixed in 10% neutral buffered formalin.

CONTROL
Two positive (for glycogen) controls + two test sections. Treat one control + one test section with diastase. One control section + one test section are NOT treated.

REAGENTS
(A) 3% ACETIC ACID
(B) ALCIAN BLUE SOLUTION (See 6.4)
(C) AMYLASE SOLUTION (See 6.2) 0.25g in 10mL distilled water . Pre-warm water for 5 sec in microwave oven then add amylase.
NOTE: AQIS requirements for disposal.
Place waste amylase into container, cap and place into yellow biohazard bag for disposal by incineration..
(D) 1% PERIODIC ACID (See 6.1)
(E) SCHIFFS REAGENT (See 6.1)

PROCEDURE
1. Deparaffinize and hydrate to distilled water.
2. Treat one control and one test section with amylase for 25 minutes.
3. Wash in running water for 20 minutes.
4. Acetic acid 2 minutes.
5. Drain.
6. Alcian blue 10 minutes.
7. Wash in tap water then distilled water.
8. Treat with 1% periodic acid for 5 minutes.
9. Rinse in distilled water.
10. Treat with Schiff's reagent 5 minutes.
11. Wash in running tap water 5-10 minutes.
12. Stain nuclei with Mayer's Haematoxylin 1 minute 30 seconds ( or 1 container of Harris haematoxylin in linear stainer).
13. Wash in water.
14. Dehydrate, clear and mount.

RESULTS
Acid mucins Blue
Neutral mucins Magenta

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6.6 BEST'S CARMINE TECHNIQUE FOR GLYCOGEN
(Best, 1906)

PRINCIPLE
This staining technique demonstrates glycogen by hydrogen bond formation between OH groups on the glycogen, and H atoms of the carminic acid. Fibrin and neutral mucin stain weakly with this method.

REFERENCES
p 219 Cook H C , Manual of Histological Demonstration Techniques p 190 Bancroft JD & Stevens A, Theory and Practice of Histological Techniques 2nd ed. SPECIMEN Standard paraffin section fixed in 10% neutral buffered formalin.

CONTROL
Liver.

REAGENTS
(A) CARMINE STOCK 60 mL H20 + 2g carmine (CI 75470) 1 g Potassium carbonate 5 g Potassium chloride Boil gently in a large conical flask. 5 mins. Cool and add 20mLs concentrated ammonia Filter and store in a dark container at 4øC. This will keep 1 - 2 months.
(B) WORKING SOLUTION Stock solution 15 mL Ammonia concentrated solution 12.5 mL Methanol 12.5 mL Staining time to be increased as stock solution ages.
(C) BEST'S DIFFERENTIATOR Methanol 40 mL Ethanol 80 mL Distilled water 100 mL PROCEDURE 1. Dewax test + positive control sections and hydrate to water. 2. Duplicate sections may be treated with diastase if desired (See6.2). 3. Stain nuclei of all sections well using one of the iron haematoxylin solutions eg. Weigerts iron haematoxylin (See 2.4) 4. Treat with carmine solution for 10 mins. 5. 4. Transfer slides quickly to a Coplin jar of Differentiating solution. 6. Wash in alcohol (NOT water), clear in histoclear and mount.

RESULTS
Glycogen Bright red Neutral mucin, mast cells, fibrin Paler red Nuclei Blue

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6.7 SOUTHGATE'S MUCICARMINE
for Epithelial Mucin This method is useful to stain encapsulated fungi of Cryptococcus neoformans, but the combined Alcian Blue - PAS technique is more informative to establish presence or absence of tissue mucins.

PRINCIPLE
Mucicarmine has large molecular size which allows the dye to penetrate and bind to acidic substrates of low density, i.e. mucins. Neutral mucins and some strongly acidic sulphated mucins do not show appreciable staining.

REFERENCE

SPECIMEN
Standard paraffin section fixed in 10% neutral buffered formalin.

REAGENTS
(A) MUCICARMINE SOLUTION Carmine 2 g Aluminium hydroxide 2 g 50% alcohol 100 mL Aluminium chloride, anhydrous 1 g Mix well, heat rapidly to boiling point and boil for 2« minutes. Cool and filter. Add the carmine and aluminium hydroxide to the alcohol in a long-necked 500 mL flask. Ground the aluminium chloride in a mortar and add it to the mixture. Note the warning. Warning: Aluminium chloride gives off HCl fumes from the bottle. Prepare in biochemistry fume hood. Wear gloves, safety glasses and mask when removing lid from and using aluminium chloride.
(B) 0.5% ACID ALCOHOL ( 0.5mL HCl in 100mL 70% alcohol)

PROCEDURE
1. Deparaffinise and hydrate to distilled water.
2. Stain with haematoxylin in linear stainer.
3. Rinse in running water.
4. Differentiate quickly in 0.5% acid alcohol. All background should be clear with only nuclei stained.
5. Rinse in running water 5 minutes.
6. Stain in Mucicarmine Solution for 2 hours.
7. Rinse in distilled water.
8. Dehydrate, clear and mount.

RESULTS
Epithelial mucin Red
Nuclei Blue

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