Lillie's Long PAS

for 1-2-Glycols

steps

materials

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The periodic acid Schiff reaction (PAS) is used to demonstrate the presence of 1-2-glycols, and is consequently an important method in the histochemistry of carbohydrates and the histological demonstration of many structures.

Lillie referred to this as

“The Periodic acid Leucofuchsin Method in Relation to Various Modifying Procedures”.

He describes its purpose as being

“To indicate at what points and in what sequence various modifying procedures should be introduced”.

Materials

A periodic acid solution.
A strong Schiff’s reagent.
Sodium metabisulfite, 0.5% aqueous, optional.
Mayer’s hemalum, or Weigert’s iron hematoxylin.
Picric acid, saturated aqueous, optional.
Orange G, 2% aqueous, optional.
Methyl blue, 0.1% in saturated aqueous picric acid, optional.

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory, although glutaraldehyde should be avoided.

Protocol

Dewax paraffin sections with xylene.
Remove xylene with two changes of absolute ethanol.

Optionally, do one or more of the following procedures in the order given.

Procedure	Comments
Acetylation	Acetylation blocks hydroxyl and amino groups. Cartilage and glycogen are the most resistant. Other carbohydrates are more easily affected.
Benzoylation	Similar to acetylation, this also blocks hydroxyl and amino groups. Again, cartilage and glycogen are the most resistant while other carbohydrates are more easily affected.
Deacetylation (saponification)	Deacetylation (also called saponification) reverses the effects of acetylation in some, if not all of the materials which were blocked. The reversal is progressive, that is, some substances recover their stainabilty before others.
Methylation	Extended methylation causes some otherwise oxidisable carbohydrates to become non-reactive. Primarily, however, it inhibits basophilia.
Aldehyde block	If applied before periodic acid oxidation, pre-existing aldehydes are inhibited from reacting and will be unstained. If applied after periodic acid oxidation, aldehydes produced by the periodic acid are inhibited from reacting and will be unstained. A second section which has not been blocked can be used to prove that oxidation formed the aldehydes.
Amylase digestion	Amylase (diastase) is used to remove glycogen. This is either to prove that a subtance is glycogen, or to remove it so that other PAS positive materials other than glycogen may be more clearly seen.
Protease digestion	Proteases may also be used to remove carbohydrate-protein complexes, although this is less common than amylase digestion because the enzymes also digest other tissue proteins.
Reducing rinses	This is an acidified solution of potassium iodide and sodium thiosulphate applied following periodic acid to remove any iodate or periodate remaining in the tissue.
Methanol, Pyridine & Solvent extraction	PAS positive carbohydrate-lipid complexes, or glycolipids, may be removed with these solvents. They may be used at ambient or elevated temperature.

Ensure sections are in water or an appropriate concentration of ethanol.
Oxidize for the appropriate time in the selected periodic acid solution.
Wash five minutes in running water, or 70% ethanol if using an alcoholic variant.
Optionally, do one of the following.
1. Hotchkiss’ reducing rinse, or
2. Aldehyde block
Place in Schiff’s reagent for 10 minutes, or longer if specially required.
Transfer to three successive sulfite rinses, 2 minutes each.
Wash in running tap water for 10 minutes.
Optionally, stain nuclei.
Optionally, stain cytoplasm.
Dehydrate and differentiate with two changes each of 95% and absolute ethanol.
Clear with xylene and coverslip using a resinous medium.

Expected Results

1-2-glycols – red
Nuclei – blue or black
Cytoplasm – yellow or unstained

Notes

A strong Schiff’s reagent is recommended, i.e. one that contains 0.5 or 1 gram of pararosanilin per 200 mL solution.
Sections should be celloidinised before deacetylation, and it may be advisable after the other options in step 3.
Experience shows that the reducing rinses may be omitted.
Mayer’s hemalum or Weigert’s iron hematoxylin are suitable nuclear stains.
Sat. picric acid, 2% orange G, or picro-methyl blue are suitable cytoplasmic stains.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

Lillie, R.D., (1954)
Histopathologic technique and practical histochemistry Ed.2
Blakiston, New York, USA.

March 30, 2023

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Lillie’s Short PAS Reaction for 1-2 Glycols

By Carmen WongAldehydes, Periodic Acid-Schiff Reaction, Protocols, Schiff's Reagent Reactions, Stain Target, Stain Type

Protocol

Lillie's Short PAS Reaction

for 1-2 Glycols

steps

materials

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Lillie referred to this method as

The Periodic acid, Schiff Sulfite Leucofuchsin Reaction, short variant.

Materials

Periodic acid (potassium iodate, nitric acid specified).
Schiff’s reagent.
Mayer’s hemalum, or Weigert’s iron hematoxylin and saturated aqueous picric acid.
Sodium metabisulfite, 0.5% aqueous
Pal’s bleach

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory, although glutaraldehyde should be avoided.

Protocol

Bring sections to water via xylene and ethanol.
Oxidize in periodic acid for ten minutes.
Wash in running water for five minutes.
Place in Schiff’s reagent for 10 minutes.
Transfer to three successive baths of sodium metabisulfite for 1, 2, and 2 minutes respectively.
Wash in running tap water for 5 minutes.
Counterstain with one of the following:
1. Place in Mayer’s hemalum for 2 minutes. Wash in running tap water and blue.
2. Place in Weigert’s iron hematoxylin for 2-4 minutes. Decolorise with Pal’s bleach diluted 1:5 with distilled water. Wash in running tap water for 4 minutes.
3. Place in Weigert’s iron hematoxylin for 6 minutes. Wash for 4 minutes in running tap water. Place in saturated aqueous picric acid for 1 minute.
Dehydrate with ethanol, clear with xylene, and coverslip using a resinous medium.

Expected Results

1-2-glycols – red
Nuclei – blue or black
Cytoplasm – yellow or unstained

Notes

The nuclear counterstain may obscure some positive staining. Keep the application time short.
Weigert’s solution should be allowed to ripen for an hour before use.
Weigert’s hematoxylin and Pal’s bleach combination gives highly selective nuclear staining.
Picric acid differentiates the nuclear stain as well as coloring the background.
Glutaraldehyde fixed tissues will have a non-specific positive background staining. This must be blocked before step 2.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

Lillie, R.D., (1954)
Histopathologic technique and practical histochemistry Ed.2
Blakiston, New York, USA.

March 30, 2023

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McManus’ PAS Reaction for 1-2 Glycols

By Carmen WongAldehydes, Periodic Acid-Schiff Reaction, Protocols, Schiff's Reagent Reactions, Stain Target, Stain Type

Protocol

McManus’ PAS Reaction

for 1-2 Glycols

steps

materials

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Materials

Periodic acid (0.5% aqueous specified).
Schiff’s reagent (Coleman’s specified).
Harris’ hemalum
Light green working solution (0.2% aqueous Light Green diluted 1:5 with distilled water)
Ammonia water (water with 3 drops concentrated ammonia per 100 mL)

Tissue Sample

6 µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory, although glutaraldehyde should be avoided.

Protocol

Bring sections to distilled water via xylene and ethanol.
Digest using a diastase, hyaluronidase, or sialidase procedure.
Oxidize in Periodic Acid for 5 minutes.
Rinse in distilled water.
Place in Coleman’s or another Schiff’s Reagent for 15 minutes.
Wash in running water for 10 minutes to develop the pink color.
Counterstain with one of the following:–
1. Harris’ hematoxylin for 6 minutes, then
  1. Wash in running water and transfer to 1% acid ethanol for 3-10 quick dips
  2. Transfer to 1% acid ethanol for 3-10 quick dips
  3. Wash in distilled water
  4. Dip in ammonia water to blue the sections
  5. Wash in running water for ten minutes
2. Light green working solution for 10 seconds.
Dehydrate with ethanol, clear with xylene, and coverslip using a resinous medium.

Expected Results

1-2-glycols – red
Nuclei – blue
Background – green (if light green used)

Notes

Light Green is better used when delineation of fungi is required.
Tap water and ammonia decolorize Light Green, so proceed directly to dehydration.
Glutaraldehyde fixed tissues will have a non-specific positive background staining. This must be blocked before step 2.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

McManus, J. F. A., (1946)
Stain Technology, v23, p99.

March 30, 2023

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Periodic Acid Schiff Reaction

By Carmen WongAldehydes, Periodic Acid-Schiff Reaction, Protocols, Schiff's Reagent Reactions, Stain Target, Stain Type

Protocol

Periodic Acid Schiff Reaction

steps

materials

print

Expected Results

Periodic acid Schiff staining in liver biopsy of glycogen storage disorder, 20X magnification

periodic-acid-schiff-staining-liver-biopsy-glycogen-storage-disorder-600x400

Figure 1. Periodic Acid Schiff Staining in Liver Biopsy of Glycogen Storage Disorder

Positive periodic acid Schiff (PAS) staining (dark purple) due to intracytoplasmic accumulation of glycogen in a liver biopsy of glycogen storage disorder shows uniformly distended hepatocytes with clear to pale eosinophilic cytoplasm. 20X magnification. Liver in glycogen storage disease, PAS stain by Department of Pathology, Calicut Medical College is licensed under CC BY-SA 4.0.

periodic-acid-schiff-staining-esophageal-candidiasis-600x400

Figure 2. Periodic Acid Schiff Staining of Esophageal Candidiasis

Periodic acid Schiff (PAS) staining (dark purple) demonstrates fungal morphology in esophageal candidiasis. Esophageal Candidiasis, PAS Stain by KGH is licensed under CC BY-SA 3.0.

1-2-glycols – red or dark purple
Nuclei – blue

Materials

Periodic acid – 1% aqueous
Schiff’s reagent
Mayer’s hemalum

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are usually satisfactory, although glutaraldehyde should be avoided.

Protocol

Bring sections to water via xylene and ethanol.
Place into periodic acid for 10-30 minutes.
Rinse well with tap water.
Rinse with distilled water.
Place in Schiff’s reagent for 10-30 minutes.
Wash off with distilled water.
Wash well with tap water for about 10 minutes.
Counterstain with Mayer’s hemalum for 2 minutes.
Wash well with tap water until hemalum is blued.
Dehydrate with ethanol, clear with xylene and coverslip using a resinous medium.

Notes

Glutaraldehyde fixation leaves free aldehyde groups attached to tissues, which causes an overall positive reaction. These groups may be stopped from reacting with an appropriate procedure such as the aniline-acetic aldehyde block.
The tap water wash at step 7 is necessary to develop the red color. Within limits, the longer the wash the darker the color.
Originally, it was recommended that the Schiff’s reagent be washed off with dilute sulfurous acid (the sulfite rinses). Since water recolors Schiff’s reagent, it was believed that a water wash could lead to false positive results. It is now known this is not the case, provided the Schiff’s reagent is removed quickly and the sections do not stay in water contaminated with it for extended periods.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

Culling C.F.A., (1974)
Handbook of histopathological and histochemical techniques Ed. 3
Butterworth, London, UK.

March 30, 2023

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Llewellyn’s PATS (Periodic acid, Thiosemicarbazide, Schmorl)

By Carmen WongAldehydes, Protocols, Stain Target

Protocol

Llewellyn's PATS

(Periodic acid, Thiosemicarbazide, Schmorl)

steps

materials

print

This technique demonstrates carbohydrates in a manner analogous to that of the periodic acid Schiff reaction, but gives a blue product.

Materials

Periodic acid, 1% aqueous.
Thiosemicarbazide, 1% aqueous
Schmorl’s solution
Material Amount
Ferric chloride, 1% aqueous, fresh 30 mL
Potassium ferricyanide, 1% aqueous, fresh 4 mL
Distilled water 6 mL
This solution should be used fresh. Prepare immediately before use.

Material	Amount
Ferric chloride, 1% aqueous, fresh	30	mL
Potassium ferricyanide, 1% aqueous, fresh	4	mL
Distilled water	6	mL

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory.

Protocol

Bring sections to water via xylene and ethanol.
Oxidize in 1% periodic acid for 10 minutes or longer.
Rinse well with water.
Place into 1% thiosemicarbazide for 5 minutes.
Wash well with running tap water to remove all traces of thiosemicarbazide.
Place into freshly made Schmorl’s solution for 10 minutes.
Wash well with water.
Counterstain with nuclear fast red.
Rinse well with water.
Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

Oxidizable carbohydrates – blue
Reducing substances – blue
Nuclei – red

Human kidney stained with periodic acid, thiosemicarbazide, Schmorl's reaction 400x magnification.

Human kidney stained with periodic acid, thiosemicarbazide, Schmorl’s reaction 400x magnification.

Human liver stained with periodic acid, thiosemicarbazide, Schmorl’s reaction 400x magnification.

Human intestine stained with periodic acid, thiosemicarbazide, Schmorl’s reaction 100x magnification.

Human intestine stained with periodic acid, thiosemicarbazide, Schmorl’s reaction 400x magnification.

Notes

It is well known that metallic azides can be explosive. However, thiosemicarbazide is not a simple metallic azide, it is a carbazide (-C=N-) and is not explosive. It is safe to use.
Reducing substances which may be present are also coloured blue. This includes melanin and enterochromaffin. Others may also be seen.
If acid hydrolysis is used instead of periodic acid oxidation (as in Feulgen’s nucleal reaction), nuclei are coloured blue.
The Schmorl’s solution is from Lillie’s modification of Schmorl’s ferricyanide reduction method for tissue reducing substances.
There is a modification of this technique for fungi.
Thiosemicarbazide has a hydrazine group at one end of it’s molecule and a thiocarbamyl group at the other. The hydrazine group combines with any aldehydes generated by periodic acid oxidation, and in so doing attaches the thiocarbamyl group to the carbohydrate. The thiocarbamyl group is a more powerful reducing group than aldehydes and rapidly reduces ferricyanide to ferrocyanide, which is immediately trapped by the ferric salt to form prussian blue at the site.
Thiosemicarbazide: (hydrazine–thiocarbamyl) = H₂NNH–CSNH₂

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

Hayashi, I., Tome, Y. and Shimosato, Y., 1989
Thiosemicarbazide used after periodic acid makes methenamine silver staining of renal glomerular basement membranes faster and cleaner.
Stain Technology, v 64, p 185.
Lillie, R.D., (1954)
Histopathologic technique and practical histochemistry Ed.2
Blakiston, New York, USA.
Llewellyn, B. D., (2014)
Thiosemicarbazide-ferricyanide reduction for the histochemical demonstration of aldehydes in tissue sections.
Biotechnic & Histochemistry, v 89, p 228-31.

March 30, 2023

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Gomori’s Methenamine Silver for Glycogen and Fungi

By Carmen WongAldehydes, Carbohydrates, Metal Impregnation, Metal Impregnation, Silver, Protocols, Stain Target, Stain Type

Protocol

Gomori's Methenamine Silver

for Glycogen and Fungi

steps

materials

print

This method is also known as Grocott’s or Grocott-Gomori’s methenamine silver.

Materials

Chromium trioxide, 5% aqu.
Neutral red, 1% aqu. or Light green SFy, 0.2% in 0.2% acetic acid, or Progressive hemalum and eosin
Sodium bisulfite, 1% aqu.
Sodium thiosulfate, 2% aqu.
Yellow gold chloride, 0.1% aqu.
Stock Methenamine silver
Material Amount
Methenamine, 3% aqu. 100 mL
Silver nitrate, 5% aqu. 5 mL
Shake until the precipitate redissolves. Silvering of the container indicates deterioration.
Working Methenamine silver
Material Amount
Stock Methenamine silver 25 mL
Distilled water 25 mL
Borax* 0.1 g
Make just before use and preheat to 50°C.
*Or 2 mL of a 5% aqueous borax solution (Grocott)

Material	Amount
Methenamine, 3% aqu.	100	mL
Silver nitrate, 5% aqu.	5	mL

Material	Amount
Stock Methenamine silver	25	mL
Distilled water	25	mL
Borax*	0.1	g

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory. A section adhesive is recommended.

Protocol

Bring sections to water via xylene and ethanol.
Oxidise with 5% chromic acid (chromium trioxide) for 60-90 minutes.
Rinse well with tap water.
Bleach with sodium bisulphite for 1 minute.
Rinse well with tap water.
Rinse with distilled water.
Treat with methenamine silver solution at 50&degC. until impregnated (up to 3 hours)
Wash with distilled water.
Tone with 0.1% gold chloride solution for 5 minutes.
Rinse with distilled water.
Fix in 2% sodium thiosulphate for 5 minutes.
Wash well with running tap water.
Counterstain with light green, neutral red or a light H&E.
Rinse with tap water.
Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

Oxidisable carbohydrates, including glycogen and fungi – black
Background – as counterstained

Notes

Methenamine is also known as hexamethylenetetramine and hexamine.
Borax is sodium tetraborate. Grocott’s modification adds the 0.1 g of borax as 2 mL of a 5% aqueous solution.
Aqueous solutions of chromium trioxide are usually referred to as chromic acid. Ten minutes in a 10% aqueous solution will usually give the same result as 60 minutes in a 5% solution.
Toning is a variable step. Untoned sections give dark brown material on a paler brown background. Many microscopists prefer to tone for about 15 seconds to produce brown-black material on a pale grey-brown background. Others tone longer (a few minutes) to produce black material on a grey background. Longer toning produces purple tones. Tone according to the personal preference of the microscopist reviewing the slides.
This method depends on a similar principle to Bauer’s chromic acid Schiff method, but in which the aldehydes produced by oxidation reduce a silver solution instead of combining with Schiff’s reagent to form a red compound. Consequently, those materials which are red in a Bauer’s stain will be black in Gomori’ stain, i.e. it is not specific for glycogen but will demonstrate any carbohydrates which can be oxidised to aldehydes, including fungi, and itis often used for that purpose.
In a similar method Hayashi, Tome and Shimosato recommended that thiosemicarbazide should be applied to the section after oxidation. Thiosemicarbazide has the formula H₂NNHCSNH₂. The hydrazine group (H₂NNH-) combines with any aldehydes present. The thiocarbamyl group (-CSNH₂) is a more powerful reducing agent than the aldehydes it replaces and reduces the methenamine silver solution more rapidly and with higher contrast.Immediately following step 5:
- Place sections in 1% aqueous thiosemicarbazide for 10 minutes.
- Wash well with tap water, and carry on from step 6.
It is well known that metallic azides can be explosive. However, thiosemicarbazide is not a simple metallic azide. The MSDS says:
- Flash Point: n/a
- Lower Explosive Limit: n/a
- Upper Explosive Limit: n/a
- Unusal Fire and Expl.rds: none identified

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

Gray, Peter. (1954)
The Microtomist’s Formulary and Guide.
Originally published by The Blakiston Co.
Republished by Robert E. Krieger Publishing Co.
Drury, R A, and Wallington, E A, (1967).
Carleton’s histological technique., Ed. 5.
Oxford University Press, London, England.
Hayashi, I., Tome, Y. and Shimosato, Y., (1989)
Thiosemicarbazide used after periodic acid makes methenamine silver staining of renal glomerular basement membranes faster and cleaner.
Stain Technology, v 64, p 185.

March 30, 2023

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Lendrum’s Chromotrope 2R for Eosinophils

By Carmen WongEosinophils, Intracytoplasmic Granules, Protocols, Stain Target

Protocol

Lendrum's Chromotrope 2R

for Eosinophils

steps

materials

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Materials

Mayer’s hemalum or similar
Carbol Chromotrope
Material Amount
Chromotrope 2R 0.5 g
Phenol 1 g
Distilled water 100 mL

Material	Amount
Chromotrope 2R	0.5	g
Phenol	1	g
Distilled water	100	mL

Preparation

Place the phenol in an Erlenmeyer flask and melt it under hot water through the glass.
Add the chromotrope 2R and mix well into a sludge.
Add the water and mix well.
Filter before use.

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory.

Protocol

Bring sections to water via xylene and ethanol.
Stain nuclei with Mayer’s hemalum and blue.
Place in the staining solution for 30 minutes.
Wash well with tap water.
Dehydrate with ethanol.
Clear with xylene and mount with a resinous medium.

Expected Results

Nuclei – blue
Eosinophil granules – bright red
Erythrocytes – may be lightly stained
Paneth cell granules – may be brownish
Enterocromaffin granules – may be brownish

Notes

The basis of this method is difficult to rationalise. Phenol is acidic and thus lowers the pH. This is often used as the basis for explaining the method, but usually an acid dye stains all basic components of the tissue (muscle, cytoplasm, collagen) intensely when applied at an acid pH. With this method eosinophils are intensely stained, but other components that would usually stain with an acidified acid dye are not. Phenol can have an intensifying effect, as is clear from its inclusion in carbol fuchsin, when it intensifies staining with basic fuchsin, but the mechanism has not been satisfactorily explained.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

Histological demonstration techniques, (1974))
Cook, H C.
Butterworths, London, England.

March 30, 2023

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del Carpio’s Impregnation for Reticulin on Previously Stained Sections

By Carmen WongMetal Impregnation, Metal Impregnation, Silver, Protocols, Reticulin, Stain Target, Stain Type

Protocol

del Carpio's Impregnation

for Reticulin on Previously Stained Sections

steps

materials

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Materials

Potassium permanganate, 0.25% aqu.
Oxalic acid, 1% aqu.
Silver nitrate, 2% aqu.
Silver nitrate, 10% aqu.
Sodium hydroxide, 40% aqu.
Ammonia, 20% aqu. (see note)
Formalin, 10% aqu.
Yellow gold chloride, 0.2% aqu.
Sodium thiosulphate, 5% aqu.
Neutral red, 1% aqu.
Ammoniacal silver

Solution Preparation

Place 1 mL of 10% silver nitrate in a flask.
Add 0.3 mL of 40% sodium hydroxide.
While swirling, slowly add drops of 20% ammonia until the precipitate just redissolves.
Make up to 100 mL with distilled water.

Tissue Sample

5 µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory. A section adhesive is recommended. The method was designed to demonstrate reticulin in sections which have been previously stained with dyes.

Protocol

Remove coverslip and bring slides to water with appropriate reagents.
Oxidise with 0.25% potassium permanganate for 20 minutes.
Rinse with tap water.
Place in Oxalic acid until bleached.
Wash with tap water.
Sensitise with 2% silver nitrate 24 hrs.
Rinse with distilled water.
Treat with ammoniacal silver for 30 minutes.
Rinse with distilled water, two changes.
Reduce in 10% formalin for 15 secs.
Rinse well with tap water.
Rinse well with distilled water.
Tone with 0.2% gold chloride solution.
Rinse with distilled water.
Fix in 5% sodium thiosulphate for 5 minutes.
Wash well with running tap water.
Counterstain with neutral red for 1 minute.
Rinse with tap water.
Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

Reticulin fibres – black
Nuclei – red
Background – grey

Notes

Strong ammonium hydroxide (s.g. 0.88) contains about 30% ammonia by weight. The instructions for making the ammoniacal silver solution specify a 20% ammonia solution, so strong ammonium hydroxide should be diluted about 2:1 with distilled water. Since the final step is to make up to 100 mL with distilled water, the precise concentration is not too important.
10% formalin is made by diluting strong formalin 1:10 with tap water (10 mL strong formalin, 90 mL tap water).
Toning is a variable step. Untoned sections give dark brown reticulin fibres on a paler brown background. Many microscopists prefer to tone for about 15 seconds to produce brown-black reticulin fibres on a pale grey-brown background. Others tone longer (a few minutes) to produce black reticulin fibres on a grey background. Longer toning produces purple tones. Tone according to the personal preference of the microscopist reviewing the slides.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

Gray, Peter. (1954)
The Microtomist’s Formulary and Guide.
Originally published by: The Blakiston Co.
Republished by: Robert E. Krieger Publishing Co.

March 30, 2023

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Simple Metachromatic Stain

By Carmen WongIntracytoplasmic Granules, Mast Cells, Protocols, Stain Target

Protocol

Simple Metachromatic Stain

steps

materials

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Materials

A metachromatic dye, 0.2% aqueous
Acetic acid, 1% aqueous
DyeSuitable blue dyes are:
Less common dyes are:

Tissue Sample

Paraffin sections of neutral buffered formalin fixed tissue are suitable. Mercuric chloride fixatives are reputed to emphasise metachromasia. Other fixatives may be satisfactory.

Protocol

Bring sections to water via xylene and ethanol.
Place in to staining solution for 5 minutes.
Rinse well with tap water.
If staining is too dark, differentiate with acetic acid until metachromasia is evident.
Rinse well with tap water.
Coverslip with water and examine wet, or blot dry and immerse in xylene. Repeat until cleared. Coverslip using a resinous medium

Expected Results

Dye color	Blue	Red	Brown
Nuclei	Blue	Red	Brown
Acid mucopolysaccharides	Red/purple	Yellow	Yellow
Background	Blue	Red	Brown

Notes

The actual dye concentration is not too important. It should be strong enough to stain within 5 minutes. Concentrations between 0.1% and 1% are usually suitable.
The staining time can be varied. The staining solution should be applied for long enough to give dark staining. Usually this is about 2-5 minutes.
Methylene blue may be polychromed by making a 1% w/v solution and leaving it for several months in an airy, bright location with a loose stopper of cotton wool. When it gives good metachromatic staining, stopper tightly and place in a cupboard.
The acetic acid concentration may be varied. Usually a concentration of 0.1% to 1% is suitable. The higher the concentration, the faster dye is removed.
Sections may require little or no differentiation. Always check the staining before applying acetic acid. Stop when a distinct contrast in color is seen.
To mount in a resinous medium, blot the wet section dry and immerse in xylene. Repeat until the section is cleared and becomes transparent. Mount using a resinous medium.
Ethanol should be avoided as it may destroy any metachromasia.
Metachromatically stained materials include intestinal and other mucins, cartilage, connective tissue ground substance and mast cell granules.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

March 30, 2023

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Cason’s Trichrome for Muscle and Collagen

By Carmen WongProtocols, Stain Type, Trichrome Staining, Trichrome, One-Step

Protocol

Cason's Trichrome

for Muscle and Collagen

steps

materials

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Materials

Staining solution

Material	Amount
Orange G	1	g
Acid fuchsin	1.5	g
Aniline blue	0.5	g
Phosphotungstic acid	1	g
Distilled water	100	mL

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory. Most trichrome stains benefit from picric acid or mercuric chloride fixation. Formalin fixed tissues may benefit from secondary fixation of sections in Bouin’s fluid.

Protocol

Bring sections to water via xylene and ethanol.
Place into the staining solution for 5 minute.
Rinse with distilled water.
Dehydrate with ethanol.
Clear with xylene and mount with a resinous medium.

Expected Results

Nuclei – red
Erythrocytes – orange
Cytoplasm – red
Collagen – blue

Notes

Although the method does not specifically call for one, an acid resistant nuclear stain could be inserted immediately before step 2.

Safety Note

Prior to handling any chemical, consult the Safety Data Sheet (SDS) for proper handling and safety precautions.

References

Gray, Peter. (1954)
The Microtomist’s Formulary and Guide.
Originally published by: The Blakiston Co.
Republished by: Robert E. Krieger Publishing Co.
Citing:
Cason, (1950)
Stain technology, vol.25, pp.225

March 30, 2023

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