Mollier’s trichrome for Elastic and Collagen

By Carmen WongDye Type, Elastic Fibers, Orcein, Protocols, Stain Target, Stain Type, Trichrome Staining, Trichrome, Multi-Step

Protocol

Mollier's trichrome

for Elastic and Collagen

steps

materials

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Materials

Weigert’s iron hematoxylin or equivalent
Solution A
Material Amount
Orcein 0.8 g
Hydrochloric acid 1 mL
Ethanol, 100% 50 mL
Distilled water 50 mL
Solution B
Material Amount
Azocarmine 2 g
Acetic acid, glacial 1 mL
Distilled water 100 mL
Solution C
Material Amount
Phosphotungstic acid 5 g
Distilled water 100 mL
Solution D
Material Amount
Naphthol green B 1 g
Acetic acid, glacial 1 mL
Distilled water 100 mL

Material	Amount
Orcein	0.8	g
Hydrochloric acid	1	mL
Ethanol, 100%	50	mL
Distilled water	50	mL

Material	Amount
Azocarmine	2	g
Acetic acid, glacial	1	mL
Distilled water	100	mL

Material	Amount
Phosphotungstic acid	5	g
Distilled water	100	mL

Material	Amount
Naphthol green B	1	g
Acetic acid, glacial	1	mL
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 solution A for 12 hours.
Stain nuclei with Weigert’s iron hematoxylin for 1-3 minutes.
Differentiate the nuclear stain if necessary.
Wash with water for 15 minutes.
Place into solution B for 15-30 minutes.
Rinse with distilled water.
decolorise in solution C for 2-6 hours, changing the solution three times.
Rinse quickly with distilled water.
Place into solution D for 15-30 minutes.
Agitate vigorously in 95% ethanol for 30 seconds.
Dehydrate with ethanol.
Clear with xylene and mount with a resinous medium.

Expected Results

Nuclei – blue
Elastic – black
Erythrocytes – red
Cytoplasm – purple
Collagen – green

Notes

Solution A is the Unna-Taenzer elastic solution.

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:
Mollier, (1938)
Zeitschrift für wissenschaftliche Mikroskopie und für mikroskopische Technik, v.55, pp.472.
And:
von Kahlden, C. and Laurent, O., (1896)
Technique microscopique, pp.143
Carré, Paris, France

March 30, 2023

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Roque’s Stain for Cell Inclusions

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

Protocol

Roque's Stain

for Cell Inclusions

steps

materials

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Materials

Stock solution A

Material	Amount
Citrate buffer, pH 5.8	100	mL
Methyl green, purified	0.1	g
Thionin	0.0165	g

Dissolve the thionin in a small amount of water. Add the buffer and methyl green. Shake and filter. Use fresh.

Citrate buffer, pH 5.8

Material	Amount
Hydrochloric acid, 0.01M	42	mL
Sodium citrate, 0.01M	58	mL

Dehydrant

Material	Amount
Tertiary butanol	80	mL
Ethanol, absolute	20	mL

Tissue Sample

Fix 2mm thick pieces of tissue in 10% formalin containing 1% sodium acetate for 3 hours. Fix smears with methanol.

Protocol

Bring sections to water via xylene and ethanol.
Place into the staining solution for 30 minutes at 40°C.
Rinse briefly with distilled water.
Place into dehydrant for 30 seconds
Replace dehydrant, 2 changes, 3 minutes each.
Rinse with absolute ethanol.
Clear with xylene and mount with a resinous medium.

Expected Results

Nuclei – blue-green
Nucleolar and cytoplasmic basophil substances – red-purple

Notes

The reference specifies methyl green, but gives the CI number for ethyl green.
The methyl green should be purified, but as a powder rather than as a solution:
- Add about 10g methyl green to 200 mL chloroform in an Erlenmeyer flask.
- Shake well.
- Filter under vacuum and in a fume chamber.
- Repeat until the chloroform is blue-green instead of violet.

Safety Note

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

References

Humason, G. L., (1967).
Animal Tissue Techniques., pp. 278
W. H. Freeman and Company, San Francisco, CA, USA.

March 30, 2023

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Bauer Reaction for Carbohydrates

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

Protocol

Bauer Reaction

for Carbohydrates

steps

materials

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Materials

A Schiff reagent
A progressive hemalum, such as Mayer
Chromic acid
Material Amount
Chromium trioxide 4 g
Distilled water 100 mL
Sulfurous acid
Material Amount
Sodium metabisulfite, 10% aqu. 6 mL
Hydrochloric acid, 1N 5 mL
Distilled water 100 mL

Material	Amount
Chromium trioxide	4	g
Distilled water	100	mL

Material	Amount
Sodium metabisulfite, 10% aqu.	6	mL
Hydrochloric acid, 1N	5	mL
Distilled water	100	mL

Tissue Sample

Protocol

Bring sections to water via xylene and ethanol.
Oxidize in chromic acid for 40-60 minutes.
Rinse with tap, then distilled water.
Place into Schiff’s reagent for 15 minutes.
Place into sulfurous acid rinses, 3 changes of 2 minutes each.
Wash with running tap water.
Counterstain with hemalum for 1 minute, and blue
Dehydrate with ethanols.
Clear with xylene and mount with a resinous medium.

Expected Results

Glycogen, mucin – red
Fungi – red
Nuclei – blue

Notes

Modern practice is to leave out the sulfite rinses and wash with large amounts of tap water.
A progressive hemalum should be used as counterstain because regressive hemalums sometimes stain mucin.
Mucins are not usually as dark as with a PAS.
Applying chromic acid for too long weakens staining due to continued oxidation of the aldehydes first produced.

Safety Note

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

References

McManus, J. F. A. and Mowry, R. W., (1960)
Staining Methods Histologic and Histochemical
Harper & Row, New York, NY, USA.

March 30, 2023

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Bensley & Bensley’s Impregnation for Reticulin

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

Protocol

Bensley & Bensley's Impregnation

for Reticulin

steps

materials

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Materials

Solutions

Material	Var I	Var II
Lugol’s iodine	+	+
Potassium permanganate, 1% aqu.	+	+
Oxalic acid, 5% aqu.	+	+
Sodium thiosulphate, 3% aqu.	+	+
Tannic acid, saturated in ethanol, 95%.	+	–
Strong ammonium hydroxide (s.g. 0.88)	+	+
Ammoniated water	+	–
Silver nitrate, aqu.	1%	2%
Formalin, 20% aqu.	+	+
Yellow gold chloride, 0.2% aqu.	+	+
Sodium hydroxide, 40% aqu.	+	+
Neutral red, 1% aqu.	+	+

Lugol’s iodine

Material	Amount
Iodine	1	g
Potassium iodide	2	g
Distilled water	300	mL

Mix the iodine and potassium iodide in a 500 mL flask. Add 5 mL of the water. When the iodine has dissolved make up to 300 mL with distilled water.

Ammoniacal silver – Var I

Place 20 mL of 1% silver nitrate in a flask. Add 4 drops of 40% sodium hydroxide Add ammonium hydroxide by drops until the precipitate is almost dissolved. Dilute 1:10 with distilled water.

Ammoniacal silver – Var I

Place 20 mL of 2% silver nitrate in a flask. Add 3 drops of 40% sodium hydroxide Add ammonium hydroxide by drops until the precipitate is just dissolved.

Tissue Sample

Bensley & Bensley said that sections of tissue fixed in Zenker, Helly, ethanol or formalin are suitable. Although they commented that Var I gave a complete impregnation of paraffin or celloidin embedded tissue, they recommended Var II for paraffin sections because of their tendency to detach from slides. A section adhesive is recommended.

Protocol

Bring sections to water via xylene and ethanol.
Oxidise in 1% potassium permanganate for 5 min.
Bleach with 5% oxalic acid.
Place in Lugol’s iodine.
Bleach in 3% sodium thiosulphate.
Wash with water.
For Var I:
1. Place in tannic acid solution at 56°C for 5 min.
2. Rinse with ammoniated distilled water.
For Var II:
1. Place in 2% silver nitrate for 16 hrs.
For Var I: Ammoniacal silver solution, VAR I, at 56°C for 10 min. For Var II: Ammoniacal silver solution, VAR II, at room temperature for 30 min.
Wash with water.
Reduce with 20% formalin for 3 min.
Wash with water.
Tone with 0.2% yellow gold chloride.
Wash with water.
Place in 3% sodium thiosulphate.
Wash with water.
Place in neutral red for 1 min.
Wash with water.
Dehydrate with ethanol, clear with xylene and mount in a resinous medium

Expected Results

Reticulin fibres – black
Nuclei – as counterstained
Background – grey or as counterstained

Notes

In the method details above, several steps do not have times given, meaning that the step is required but no other details were given. Common sense should prevail, and the step done for sufficient time to accomplish its obviously intended purpose. If it is a water wash for removal of an excess of the preceding material it would usually be for approximately 1-2 minutes. If it is for toning with gold chloride then see the final note below.
The method details also often specify to “Wash in water” without saying whether distilled or tap water should be used. In many cases it does not matter, but common sense should prevail. If tap water is likely to produce a non-specific precipitate of silver, then use distilled water and, when it specifies to “wash”, give several changes. Tap water varies in quality and individual laboratory’s results may differ due to that. Of course, distilled water could be used throughout, but it is strongly recommended after the silver or gold chloride solutions since these may be affected by tap water contaminants.
Bensley & Bensley said that “the silver carbonate solution of Hortega” could be substituted for their own silver oxide solution in Var I. Hortega gave form several such solutions and the authors do not say which one they meant. These formulas differ mainly by the amounts of 10% aqueous silver nitrate added to 5% aqueous sodium carbonate. All redissolve the resulting precipitate with drops of strong ammonium hydroxide.Hortega’s Ammoniacal silver solutions
- Place 50 mL of 5% sodium carbonate in a flask. Add 12 mL of 10% silver nitrate. Let the precipitate settle, then decant the supernatent. Wash, allow to settle and decant several times. Add ammonium hydroxide by drops until the precipitate is almost dissolved. Dilute to 100 mL with distilled water.
- In addition to the formula above, another adds 12.5 mL silver nitrate, does not decant and wash, but does dilute to 100 mL with distilled water.
- A third adds 20 mL silver nitrate to 80 mL sodium carbonate, does not decant and wash, and does not dilute with distilled water.
- A fourth adds 25 mL silver nitrate to 75 mL sodium carbonate, does not decant and wash, and does not dilute with distilled water.
- A fifth adds 12 mL silver nitrate to 50 mL saturated aqueous lithium carbonate, decants and washes, and dilutes to 100 mL with distilled water.
Ensure that the ammonium hydroxide is fresh and full strength. Keep well stoppered when not in use. After removing the amount required immediately restopper the bottle.
Improperly made ammoniacal silver solutions can affect the quality of the impregnation. There should be a faint, persistent opalescence, with only a faint smell of ammonia.
20% formalin is made by diluting 20 mL strong formalin with 80 mL water.
Bensley & Bensley suggested either Heidenhain’s Azan or 1% aqueous acridine red as counterstains. I have substituted neutral red.
The formula given for Lugol’s iodine is now usually referred to as Gram’s iodine.
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

Bensley R. R. and Bensley, S. H., (1938)
Handbook of Histological and Cytological Technique.
U. Chicago Press, Chicago, USA
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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Basic Fuchsin–Picric Acid for Elastic Fibers

By Carmen WongElastic Fibers, Protocols, Stain Target

Protocol

Basic Fuchsin–Picric Acid

for Elastic Fibers

steps

materials

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Materials

Basic fuchsin

Material	Amount
Basic fuchsin	0.5	g
Distilled water	500	mL

Picric acid

Material	Amount
Saturated alcoholic picric acid	12	mL
Acetone	1	L

Tissue Sample

A 10% Formalin variant is suitable. Paraffin sections at 5µ are preferred.

Protocol

Bring sections to water via xylene and ethanol.
Place in basic fuchsin for 1-2 minutes.
Rinse with tap water, then acetone.
Differentiate briefly with picro-acetone until tissue is just decolorised.
Rinse with acetone.
Clear with xylene and mount with a resinous medium

Expected Results

Elastic fibres – red
Other tissues – yellow

Notes

The Brown and Brenn picric acetone usually requires weighing semi-dry picric acid. The amount in the saturated ethanolic solution specified above is very close to that and it is a much safer means of compounding the solution.
If you use the Brown and Brenn variant of the Gram stain, the counterstain solutions from that may be used.
Be careful not to overdifferentiate. Apply picric acetone until most of the red is just removed from the background. This takes only a few seconds. It is easy to overdifferentiate.
This method is not meant as a primary means of staining elastic fibres. It can be done very quickly, about 5-10 minutes, and may be useful when time is limited.
Nuclei are not stained.

Safety Note

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

References

Personal observation, Bryan D. Llewellyn.
Brown, J H and Brenn, L, (1931),
A method for the differential staining of Gram positive and Gram negative bacteria in tissue sections.,
Bull. John Hopkins Hosp., v 48, page 69.

March 30, 2023

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Buffered Thionin for Nissl Bodies

By Carmen WongIntracytoplasmic Granules, Nissl Bodies, Protocols, Stain Target

Protocol

Buffered Thionin

for Nissl Bodies

steps

materials

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Materials

Material	Amount for pH 3.7 Solution		Amount for pH 4.5 Solution
Acetic acid, 0.6% (0.1M)	90	mL	60	mL
Sodium acetate, 0.8% (0.1M)	10	mL	40	mL
Thionin, 1% aqueous	2.5	mL	2.5	mL

Tissue Sample

10µ paraffin sections fixed in 10% formalin variants or Carnoy’s chloroform-ethanol-acetic mixture are suitable. Other fixatives may be satisfactory.

Protocol

Standard Method

Bring sections to water via xylene and ethanol.
Place into one of the staining solutions for 20-60 minutes.
Dehydrate with ascending concentrations of ethanol.
Clear with xylene and mount with a resinous medium.

Alternative Method

Dilute the thionin with distilled water instead of acetate buffer.
Bring sections to water via xylene and ethanols.
Stain in aqueous thionin for 20-60 minutes.
Rinse with ethanol, 50%.
Differentiate with 0.25% acetic acid in 95% ethanol, controlling microscopically.
Rinse well with 95% ethanol.
Complete dehydration with absolute ethanol.
Clear with xylene and coverslip using a resinous medium.

Expected Results

Structure	pH 3.7 Staining Solution	pH 4.5 Staining Solution
Nissl bodies	blue	dark blue
Nuclei	blue	dark blue
Background	pale or unstained	pale blue

Safety Note

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

References

Davenport, H.A.. (1960).
Histological and Histochemical Technics,
W. B. Saunders, Philadelphia, USA.
Citing:
Windle, W. F., Rhines, R. and Rankin, J. (1943),
A Nissl method using buffered solutions of thionin.
Stain Technology, v 8, pp. 77-86.
and:
Conn, H. J. and Darrow, M. A.,, (1946),
Staining procedures.
Biotech Publications, Geneva, New York.

March 30, 2023

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Burns, Pennock & Stoward’s Thioflavine T for Amyloid Fluorescence

By Carmen WongAmyloid, Fluorescent Staining, Protocols, Stain Target, Stain Type

Protocol

Burns, Pennock & Stoward's Thioflavine T

for Amyloid Fluorescence

steps

materials

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Materials

Krutsay’s hemalum
Thioflavine T
Material Amount
Thioflavine T 0.5 g
Hydrochloric acid, 1N 100 mL

Material	Amount
Thioflavine T	0.5	g
Hydrochloric acid, 1N	100	mL

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable.

Protocol

Bring sections to water via xylene and ethanol.
Stain nuclei with alum hematoxylin.
Rinse well with water.
Stain with thioflavine T solution for 5 minutes.
Blot, rinse well with absolute ethanol.
Mount in a fluorescence free resinous mounting medium.

Expected Results

Using a UG1 or UG2 exciter filter and a UV barrier filter, or a BG12 exciter and an OG4 or OG5 barrier filter, amyloid fluoresces lime green or blue.

Safety Note

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

References

Burns, J., Pennock, C.A. and Stoward, P.J., (1967),
The specificity of the staining of amyloid deposits with thioflavine T,
Journal of pathology and bacteriology, v 94, page 337.
Bancroft, J.D. and Stevens A. (1982)
Theory and practice of histological techniques Ed. 2
Churchill Livingstone, Edinburgh & London, UK.

March 30, 2023

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Llewellyn’s Sirius Red for Amyloid

By Carmen WongAmyloid, Direct Dye Staining, Eosinophils, Intracytoplasmic Granules, Paneth Cells, Protocols, Stain Target, Stain Type

Protocol

Llewellyn's Sirius Red

for Amyloid

steps

materials

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Materials

Material	Amount
Sirius red F3B	0.5	g
Distilled water	50	mL
Ethanol, absolute	50	mL

Staining Solution Preparation

Dissolve the dye into the water, add ethanol and mix well.
Add 1 mL of 1% sodium hydroxide. Then, while strong backlighting and swirling, add drops of 20% sodium chloride until a fine haze is detected. Usually about 2 mL is adequate. Adding more than 4 mL causes excessive precipitation. The solution is reasonably stable for several months, but slowly deteriorates. Extend the staining time to compensate. When it requires more than 2 hours to adequately stain, prepare a new solution.

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 a progressive alum hematoxylin for a few minutes.
Rinse with tap water.
Rinse with ethanol.
Place into alkaline sirius red for 1 – 2 hours.
Rinse well with tap water.
Dehydrate with absolute ethanol.
Clear with xylene and mount with a resinous medium.

Expected Results

Amyloid – red
Eosinophil and Paneth cell granules – red
Nuclei – blue
Background – colorless

Notes

Amyloid displays deep green birefringence when viewed with crossed polarisers, one above and one below the section.
Eosinophils and Paneth cell granules are also demonstrated. If used for this purpose the sodium chloride may be ommitted.
This method uses sirius red F3B. The dye Sirius red 4B is not suitable.

Safety Note

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

References

Llewellyn, B.D., (1970)
An improved sirius red method for amyloid.
Journal of Medical Laboratory Technology, v 23, 308

March 30, 2023

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Puchtler, Sweat and Levine’s Congo Red for Amyloid

By Carmen WongAmyloid, Direct Dye Staining, Protocols, Stain Target, Stain Type

Protocol

Puchtler, Sweat and Levine's Congo Red

for Amyloid

steps

materials

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Materials

Mayer’s hemalum
Stock alcoholic salt
- Saturate 80% ethanol with sodium chloride.
Stock congo red
- Saturate 80% ethanol with congo red and sodium chloride.
- Let stand for 24 hours.
Working alkaline alcohol
Material Amount
Stock alcoholic salt 50 mL
1% sodium hydroxide 0.5 mL
Use within 15 minutes.
Working congo red
Material Amount
Stock congo red 50 mL
1% sodium hydroxide 0.5 mL
Use within 15 minutes.

Material	Amount
Stock alcoholic salt	50	mL
1% sodium hydroxide	0.5	mL

Material	Amount
Stock congo red	50	mL
1% sodium hydroxide	0.5	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.
Stain nuclei with hemalum.
Rinse well with distilled water.
Place in working alkaline alcohol for 20 minutes.
Place in working congo red for 20 minutes.
Dehydrate rapidly with absolute ethanol.
Clear with xylene and mount with a resinous medium.

Expected Results

Amyloid – Deep pink to red
Nuclei – blue
Background – colorless

Notes

Amyloid displays apple green birefringence when viewed with crossed polarizers, one above and one below the section.
This method is considered to be the most reliable of all congo red methods for amyloid.

Safety Note

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

References

Puchtler, H., Sweat, F. and Levine, M., (1962),
On the binding of congo red by amyloid,
Journal of Histochemistry and Cytochemistry, v 10, page 355
Bancroft, J.D. and Stevens A. (1982)
Theory and practice of histological techniques Ed. 2
Churchill Livingstone, Edinburgh & London, UK.

March 30, 2023

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Stokes’ Congo Red for Amyloid

By Carmen WongAmyloid, Direct Dye Staining, Protocols, Stain Target, Stain Type

Protocol

Stokes' Congo Red

for Amyloid

steps

materials

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Materials

Mayer’s hemalum
80% ethanol
Alkaline Congo Red
Material Amount
Congo red as required
Ethanol, 80% 100 mL
Potassium hydroxide 0.2 g

Material	Amount
Congo red	as	required
Ethanol, 80%	100	mL
Potassium hydroxide	0.2	g

Compounding Procedure

Dissolve the potassium hydroxide in the ethanol.
Add sufficient dye to saturate. Leave overnight and filter.
Stable for about 3 months.

Tissue Sample

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

Protocol

Bring sections to water via xylene and ethanol.
Place into alkaline congo red for 25 minutes.
Wash with tap water for 5 minutes.
Stain nuclei with Mayer’s hemalum for 5 minutes.
Blue in running tap water for 10 minutes.
Dehydrate rapidly in absolute ethanol.
Clear with xylene and mount with a resinous medium.

Expected Results

Amyloid – orange to red
Nuclei – blue
Background – colorless

Notes

If congo red is applied for longer than 25 minutes, the background will show some coloration.
Green birefringence is displayed under crossed polarizers.

Safety Note

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

References

Stokes, Gwen, (1976),
An improved congo red method for amyloid,
Medical Laboratory Sciences, v 33, 79

March 30, 2023

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