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Stain Target

Akita & Kaneko’s Hemalum for Elastic Fibers

By Dye Type, Elastic Fibers, Hematoxylin and Eosin Staining, Mordanted Hematoxylin, Protocols, Stain Target, Stain Type
Protocol

Akita & Kaneko's Hemalum

for Elastic Fibers

5
steps
6
materials
print

Materials

MaterialAmount
Hematoxylin100mg
Ethanol, 70%100mL
Sodium iodate20mg
Potassium alum300mg
Chloral hydrate5g
Citric acid100mg

Staining Solution Preparation

  1. Dissolve the hematoxylin into the ethanol.
  2. Add the potassium alum, then the other ingredients.
  3. Mix well for 15 to 20 minutes, then filter.
  4. At this point the solution is stable for 2 months at room temperature.
  5. Immediately prior to use, adjust to pH 8.0 with saturated aquueous lithium carbonate or 1N potassium hydroxide.

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Helly and Bouin are also suitable. Other fixatives should be tested.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into freshly pH adjusted staining solution for 30-60 minutes.
  3. Wash with running tap water for 10 minutes.
  4. Dehydrate with ethanols.
  5. Clear with xylene and mount with a resinous medium.

Expected Results

  • Elastic fibres  –  blue
  • Collagen  –  pale violet

Notes

  • Note that this is an ethanolic variant of Mayer’s hemalum.
  • If sodium iodate is left out, the staining solution lasts 3-4 days, but the staining time is extended to 60 minutes.
  • The solution may also be made by diluting 30 mL Mayer’s hemalum with 70 mL absolute ethanol, mixing for 20 minutes, and filtering before adjusting the pH. Staining requires 60 minutes and the results are paler.

Safety Note

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

References

  1. Akita, M. and Kaneko, K., (1981)
    An improved hematoxylin method with application of Mayer’s hemalum for staining of elastic fibres.
    Stain Technology, v. 56, p. 327
  2. Kaneko, K. and Akita, M., (1978)
    On the staining of elastic fibres with alum hematoxylin.
    Stain Technology, v. 53, p. 43.
March 30, 2023
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Lendrum, Slidders & Fraser’s Alcian Blue for amyloid

By Amyloid, Protocols, Stain Target
Protocol

Lendrum, Slidders & Fraser's Alcian Blue

for amyloid

10
steps
10
materials
print

Materials

  • Stock A
    Alcian blue 8GX1g
    Ethanol 80%100mL
  • Stock B
    Sodium sulfate, hydrate1g
    Distilled water100mL
  • Working solution
    Stock A45mL
    Stock B45mL
    Glacial acetic acid10mL

    Mix. Leave 30 minutes. Prepare daily.

  • Acetic ethanol
    Ethanol, 95%45mL
    Distilled water45mL
    Glacial acetic acid10mL

    Prepare daily.

  • Borax Ethanol
    Boraxasrequired
    Ethanol, 80%100mL

    Dissolve borax to saturation.

Tissue Sample

Paraffin sections of formalin fixed tissues are satisfactory. If using the more complex trichrome counterstains, formal sublimate fixation is preferred.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into acetic alcohol for 1 – 2 minutes.
  3. Place into the working solution for 2 hours.
  4. Place into acetic alcohol for 1 – 2 minutes.
  5. Wash with water.
  6. Place into alcoholic borax for 30 minutes.
  7. Wash with water.
  8. Counterstain (see notes).
  9. Dehydrate with absolute ethanol.
  10. Clear with xylene and mount with a resinous medium.

Expected Results

  • Amyloid  –  blue green
  • Other tissue  –  according to the counterstain

Notes

  • The simplest counterstain is the van Gieson.
  • If counterstaining with PAS, treatment with alcoholic borax may be omitted.
  • If counterstaining with silver impregnation for reticulin, the method must not include a Mallory bleach.
  • The authors suggested their own, more complex trichrome counterstain.

Safety Note

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

References

  1. Lendrum, A.C., Slidders, W. and Fraser, S., (1972),
    Renal hyalin: A study of amyloidosis and diabetic fibrinous vasculosis with new staining methods.,
    Journal of Clinical Pathology, v 25, page 373.
March 30, 2023
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Gabe’s Aldehyde Fuchsin

By Aldehyde Fuchsin, Dye Type, Elastic Fibers, Protocols, Stain Target, Stain Type
Protocol

Gabe's Aldehyde Fuchsin

6
steps
9
materials
print

Materials

Stock powder

MaterialAmount
Pararosanilin5g
Paraldehyde5mL
Hydrochloric acid10mL
Distilled water1L

Dye Preparation Procedures

  1. Add the dye to the water and boil.
  2. Cool to room temperature.
  3. Add paraldehyde and hydrochloric acid.
  4. Ripen at RT two days. Filter. Wash precipitate with 50mL distilled water. Dry the ppt and paper at 60°C.
  5. Store in a labelled container. Stable for about ten years.

Working solution

MaterialAmount
Stock powder0.25g
Ethanol, 70%200mL
Acetic acid, glacial2mL

Acid ethanol

MaterialAmount
Hydrochloric acid, conc.1mL
Ethanol, 95%200mL

Tissue Sample

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

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place in the working solution for 5 minutes.
  3. Rinse well with running tap water.
  4. Rinse with acid ethanol for 20-30 seconds to remove excess dye.
  5. Counterstain if wished.
  6. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Elastic fibres  –  purple
  • Mast cells  –  purple
  • Pituitary βcells  –  purple
  • Sulphated mucins  –  purple
  • Background  –  as the counterstain
  • Nuclei  –  as the nuclear stain

Notes

  • The basic fuchsin used for this solution should be one that is suitable for Schiff’s reagent, i.e., it should have a high pararosanilin content. Both methods involve forming a compound between an aldehyde and dye.
  • Light counterstaining with a progressive alum hematoxylin and eosin is also suitable.
  • Many other counterstains can be used, including methods such as Masson’s trichrome.

Safety Note

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

References

  1. Kiernan. J.A., (1999)
    Histological and histochemical methods: Theory and practice, Ed. 3
    Butterworth Heinemann, Oxford, UK.
    Citing:
    Gabe, M., (1976)
    Histological techniques.
    Masson, Paris.
March 30, 2023
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Gomori’s Aldehyde Fuchsin

By Aldehyde Fuchsin, Dye Type, Elastic Fibers, Intracytoplasmic Granules, Mast Cells, Protocols, Stain Target, Stain Type
Protocol

Gomori's Aldehyde Fuchsin

7
steps
4
materials
print

Materials

Solution A

MaterialAmount
Basic fuchsin1g
Paraldehyde, fresh1mL
Hydrochloric acid, conc.1mL
Ethanol, 70%200mL

Compounding Procedure

  1. Dissolve the dye in the ethanol.
  2. Add paraldehyde and hydrochloric acid.
  3. Ripen at room temperature for 48-72 hours.
  4. Refrigerate. The solution is stable for 2-3 months.

Tissue Sample

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

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Wash with water.
  3. Rinse with 70% ethanol.
  4. Place in the staining solution for 10 minutes.
  5. Rinse well with 95% ethanol.
  6. Counterstain the nuclei and/or the cytoplasm if wished.
  7. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Elastic fibres  –  purple
  • Mast cells  –  purple
  • Pituitary β cells  –  purple
  • Sulphated mucins  –  purple
  • Background  –  as the counterstain
  • Nuclei  –  as the nuclear stain

Notes

  • The basic fuchsin used for this solution should be one that is suitable for Schiff’s reagent, i.e., it should have a high pararosanilin content. Both methods involve forming a compound between an aldehyde and dye.
  • Light counterstaining with a progressive alum hematoxylin and eosin is also suitable.
  • Many other counterstains can be used, including methods such as Masson’s trichrome.
  • Gabe described a technique for the preparation and use of aldehyde fuchsin powder.

Safety Note

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

References

  1. Drury, R.A.B. and Wallington, E.A., (1980)
    Carleton’s histological technique Ed. 5
    Oxford University Press, Oxford, UK.
March 30, 2023
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Aldehyde Toluidine Blue for Mast Cells

By Intracytoplasmic Granules, Mast Cells, Protocols, Stain Target
Protocol

Aldehyde Toluidine Blue

for Mast Cells

7
steps
6
materials
print

Materials

Compounding Procedure

  1. Dissolve the dye in the ethanol.
  2. Add paraldehyde and acid.
  3. Ripen one week at room temperature.
  4. Store at room temperature.
  5. Filter before use. Stable for a year or longer.

Tissue Sample

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

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Rinse with 70% ethanol.
  3. Place in solution A for 1 hour.
  4. Wash off excess stain with 70% ethanol.
  5. Rinse well with tap water.
  6. Counterstain with nuclear fast red-tartrazine.
  7. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Mast cell granules  –  deep blue
  • Nuclei  –  red
  • Background  –  yellow

Notes

  • The staining solution is a modification of Gomori’s aldehyde fuchsin using toluidine blue instead of basic fuchsin.
  • Staining time may need to be increased as the solution ages (up to 2 hours). If staining takes longer than 2 hours, prepare a new solution.
  • Elastic fibres are unstained, likely because basic fuchsin can form dipole-dipole interactions and toluidine blue generally does not.
  • Mucins are stained very pale blue.
  • Different samples of this dye may vary in effectiveness. If a sample gives pale staining, try one from another vendor. Toluidine blue from Fisher Scientific was used to develop the method.

Safety Note

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

References

  1. Llewellyn, B. D., unpublished.
March 30, 2023
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Naphthoic Acid Hydrazide for Aldehydes

By Aldehydes, Protocols, Stain Target
Protocol

Naphthoic Acid Hydrazide

for Aldehydes

11
steps
11
materials
print

Materials

Veronal acetate

MaterialAmount
Sodium acetate1.943g
Sodium barbiturate2.943g
Distilled waterto 100mL

Veronal buffer pH 7.4

MaterialAmount
Veronal acetate solution5mL
M/10 hydrochloric acid5mL
Distilled water60mL

NAH

MaterialAmount
2-hydroxy-3-naphthoic acid hydrazide0.1g
Ethanol, 100%95mL
Acetic acid, glacial5mL

Fast blue B

MaterialAmount
Fast blue B salt0.1g
Veronal acetate buffer (pH 7.4)100mL

Tissue Sample

5 µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Many other fixatives are satisfactory. Fixatives containing strong acids should be avoided if the intent is to demonstrate aldehydes generated from acid hydrolysis of DNA, as acids in some fixatives may hydrolyse the tissue during fixation (picric acid in Bouin’s formal-picric-acetic mixture, for example).

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Oxidise or hydrolyse to generate aldehydes.
  3. Rinse briefly with distilled water.
  4. Rinse briefly with 50% ethanol.
  5. Place into NAH solution at room temperature for 3-6 hours.
  6. Rinse with three changes of 50% ethanol, about 10 minutes each.
  7. Wash well with water.
  8. Place into pre-cooled fast blue B solution for 1-3 minutes at 0°C.
  9. Wash well with water.
  10. Optionally, counterstain appropriately.
  11. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Aldehyde sites  –  blue to purple
  • Background  –  as counterstained

Notes

  • Sodium barbiturate is also known as veronal.
  • Procedures for producing aldehydes include those for acid hydrolysis of DNA, periodic acid and chromic acid oxidation of carbohydrates. In those procedures, begin at step 3, above, where those other methods specify placing into Schiff’s reagent.

Safety Note

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

References

  1. Culling, C F A, Allison, R T, Barr, W T, (1985).
    Cellular pathology technique., Ed. 4., p. 187
    Butterworths, London, England.
March 30, 2023
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Cook & Lamb Alkali Blue for Elastic Fibres

By Elastic Fibers, Protocols, Stain Target
Protocol

Cook & Lamb Alkali Blue

for Elastic Fibres

10
steps
3
materials
print

Materials

  • 0.4% alkali blue in 70% ethanol
  • 4% aqueous ferric ammonium sulphate
  • 0.05% potassium hydroxide in absolute ethanol

Tissue Sample

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

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place in alkali blue solution for 5 minutes.
  3. Wash with water.
  4. Place in iron alum solution for 3 minutes.
  5. Wash with water.
  6. Rinse with ethanol.
  7. Rinse briefly with alcoholic potassium hydroxide to clear the background.
  8. Counterstain if wished.
  9. Dehydrate with ethanols.
  10. Clear with xylene and mount with a resinous medium.

Expected Results

  • Elastic fibres – blue (blue-green after Van Gieson)
  • Erythrocytes – pale blue-green

Notes

  • Neutral red, safranin, hemalum and eosin are suggested as counterstains.
  • The alkali blue solution lasts for several weeks.
  • This is an example of afterchrome mordanting.

Safety Note

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

References

  1. Histological demonstration techniques, (1974))
    Cook, H C.
    Butterworths, London, England
March 30, 2023
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Allen’s Stain for Mast Cells

By Intracytoplasmic Granules, Mast Cells, Protocols, Stain Target
Protocol

Allen's Stain

for Mast Cells

11
steps
3
materials
print

Materials

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

  1. Bring sections to water via xylene and ethanol.
  2. Rinse with water.
  3. Stain nuclei lightly with alum hematoxylin.
  4. Rinse with tap water and blue hematoxylin.
  5. Rinse well with water.
  6. Place in neutral red for 10 minutes.
  7. Rinse with distilled water.
  8. Differentiate with 70% ethanol up to 10 minutes.
  9. Dehydrate with 96% ethanol up to 5 minutes.
  10. Dehydrate with N-butanol up to 10 minutes.
  11. Clear with xylene and mount using a resinous medium.

Expected Results

  • Nuclei – blue
  • Mast cell granules = red

Safety Note

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


References

  1. Culling, C F A, (1976).
    Lynch’s Medical Laboratory Technology., Ed. 3. Vol. II. pp. 980
    W. B. Saunders Company, Toronto, Canada.
  2. Putt, F A, (1972).
    Manual of Histopathological Staining Methods., pp. 233
    John Wiley & Sons, London, UK.
March 30, 2023
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Bennhold’s Congo Red for Amyloid

By Amyloid, Direct Dye Staining, Protocols, Stain Target, Stain Type
Protocol

Bennhold's Congo Red

for Amyloid

8
steps
5
materials
print

Materials

  • Mayer’s hemalum
  • Congo red
    MaterialAmount
    Congo red1g
    Distilled water100mL
  • Lithium carbonate
    MaterialAmount
    Lithium carbonateasrequired
    Distilled water100mL

Dissolve to saturation.

Tissue Sample

Paraffin sections of formalin fixed tissues are satisfactory.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into the congo red solution for
    1. 30 minutes at room temperature, or
    2. 45 minutes at 56°C, or
    3. 15 seconds at boiling.
  3. Drain and, without rinsing, place into lithium carbonate solution for 15 seconds.
  4. Drain and, without rinsing, differentiate in 80% ethanol. This usually takes just a few seconds.
  5. Wash well with tap water.
  6. Stain nuclei with hematoxylin and blue.
  7. Dehydrate with ethanol.
  8. Clear with xylene and mount with a resinous medium.

Expected Results

  • Amyloid  –  red
  • Nuclei  –  blue

Notes

  • Amyloid displays apple green birefringence when viewed with crossed polarisers, one above and one below the section.
  • The congo red is most commonly applied for 30 minutes at room temperature.
  • Differentiation in 80% ethanol is difficult to control, and amyloid is often poorly demonstrated. Due to this the method is not usually recommended.

Safety Note

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

References

  1. Bennhold, H., (1922),
    Eine specifische amyloidfärbung mit kongorot,
    Münchener Medizinische Wochenschrift, v 44, page 1537
March 30, 2023
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Congo Red Fluorescence for Amyloid

By Amyloid, Direct Dye Staining, Protocols, Stain Target, Stain Type

Protocol

Congo Red Fluorescence

for Amyloid

8
steps
4
materials
print

Expected Results

Congo red staining showing amyloid and lipofuscin in an autopsy specimen of senile cardiac amyloidosis

fig1-congo-red-staining-amyloid-lipofuscin-autopsy-senile-cardiac-amyloidosis-600x400

Figure 1. Congo Red Staining Showing Amyloid and Lipofuscin in an Autopsy Specimen of Senile Cardiac Amyloidosis

Congo red staining of an autopsy specimen of senile cardiac amyloidosis, showing amyloid (extracellular washed-out red material) and abundant lipofuscin (yellow granular material). Cardiac amyloidosis very high mag by Nephron / Michael Bonert, Pathology & Molecular Medicine, McMaster University is licensed under CC BY-SA 3.0.

  • Amyloid  –  Orange to red fluorescence

Materials

Congo red

MaterialAmount
Congo red0.1g
Ethanol, 50%100mL

Differentiator

MaterialAmount
Potassium hydroxide0.2g
Ethanol, 80%100mL

Tissue Sample

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

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into congo red for 1 minute.
  3. Wash with tap water.
  4. Place into differentiator until the section appears to be unstained.
  5. Wash with water.
  6. Dehydrate with absolute ethanol.
  7. Clear with xylene.
  8. Mount with a fluorescence free resinous medium.

Notes

  • This method is very similar to Highmans procedure.

Safety Note

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

References

  1. 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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