Category

Stain Type

Puchtler, Sweat and Levine’s Congo Red for Amyloid

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

Puchtler, Sweat and Levine's Congo Red

for Amyloid

7
steps
10
materials
print

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
    MaterialAmount
    Stock alcoholic salt50mL
    1% sodium hydroxide0.5mL

    Use within 15 minutes.

  • Working congo red
    MaterialAmount
    Stock congo red50mL
    1% sodium hydroxide0.5mL

    Use within 15 minutes.

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. Stain nuclei with hemalum.
  3. Rinse well with distilled water.
  4. Place in working alkaline alcohol for 20 minutes.
  5. Place in working congo red for 20 minutes.
  6. Dehydrate rapidly with absolute ethanol.
  7. 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

  1. 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
  2. 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 Amyloid, Direct Dye Staining, Protocols, Stain Target, Stain Type
Protocol

Stokes' Congo Red

for Amyloid

7
steps
5
materials
print

Materials

  • Mayer’s hemalum
  • 80% ethanol
  • Alkaline Congo Red
    MaterialAmount
    Congo redasrequired
    Ethanol, 80%100mL
    Potassium hydroxide0.2g

Compounding Procedure

  1. Dissolve the potassium hydroxide in the ethanol.
  2. Add sufficient dye to saturate. Leave overnight and filter.
  3. 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

  1. Bring sections to water via xylene and ethanol.
  2. Place into alkaline congo red for 25 minutes.
  3. Wash with tap water for 5 minutes.
  4. Stain nuclei with Mayer’s hemalum for 5 minutes.
  5. Blue in running tap water for 10 minutes.
  6. Dehydrate rapidly in absolute ethanol.
  7. 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

  1. Stokes, Gwen, (1976),
    An improved congo red method for amyloid,
    Medical Laboratory Sciences, v 33, 79
March 30, 2023
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Sweat and Puchtler’s Sirius Red for Amyloid

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

Sweat and Puchtler's Sirius Red

for Amyloid

11
steps
8
materials
print

Materials

  • Mayer’s hemalum
  • Neutral buffered formalin (pH 7.0)
  • 0.1M borate or borate-phosphate buffer pH 9.0
  • Alkaline alcohol
    MaterialAmount
    Ethanol, 80%100mL
    Sodium hydroxide, 1% aqueous1mL
  • Sirius red
    MaterialAmount
    Sirius red1g
    Distilled water100mL
    Sodium chloride0.5g

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 neutral buffered formalin overnight.
  3. Wash in tap water for 15 minutes.
  4. Place into alkaline alcohol for 20 – 60 minutes.
  5. Rinse well with distilled water.
  6. Place into sirius red at 60°C for 60 – 90 minutes.
  7. Rinse with buffer.
  8. Wash with tap water for 5 minutes.
  9. Stain nuclei with hemalum and blue.
  10. Dehydrate with absolute ethanol.
  11. Clear with xylene and mount with a resinous medium.

Expected Results

  • Amyloid  –  red
  • Nuclei  –  blue
  • Background  –  colorless

Notes

  • This method uses sirius red F3B. The dye sirius red 4B is not suitable.
  • Sirius scarlet GG, CI 40270, may also be used.
  • Amyloid displays deep green birefringence when viewed with crossed polarizers, one above and one below the section.

Safety Note

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

References

  1. Sweat, F. and Puchtler, H., (1965),
    Demonstration of amyloid with direct dyes,
    Archives of Pathology, v 80, page 613
March 30, 2023
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Thioflavine S For Senile Plaques & Neurofibrillary Tangles In Alzheimer’s

By Amyloid, Fluorescent Staining, Protocols, Stain Target, Stain Type
Protocol

Thioflavine S

For Senile Plaques & Neurofibrillary Tangles In Alzheimer's

13
steps
13
materials
print

Materials

  • Potassium permanganate, 0.25% in distilled water
  • Ethanol, 70%
  • Acetic acid, 0.25%
  • Bleach solution
    MaterialAmount
    Potassium metabisulphite1g
    Oxalic acid1g
    Distilled water100mL
  • Blocking solution
    MaterialAmount
    Sodium hydroxide1g
    Hydrogen peroxide, 30%3mL
    Distilled water100mL
  • Thioflavine S
    MaterialAmount
    Thioflavine S0.0125g
    Ethanol, 50%100mL
  • Glycerine Water
    MaterialAmount
    Glycerine3vols.
    Distilled water1vol.

Tissue Sample

30µ free floating sections of neutral buffered formalin fixed tissue are suitable. If paraffin embedded, they should be carefully dewaxed and hydrated before staining, but should remain free floating.

Protocol

  1. Place in potassium permanganate for 20 minutes.
  2. Rinse well with distilled water.
  3. Place in the bleach solution for 2 minutes.
  4. Rinse well with distilled water.
  5. Place in the blocking solution for 20 minutes.
  6. Rinse well with distilled water.
  7. Place in acetic acid solution for 5 seconds.
  8. Rinse well with distilled water.
  9. Mount sections on microscope slides using an adhesive, dry, then rehydrate.
  10. Place into thioflavine S solution for 3-5 minutes.
  11. Rinse twice with 50% ethanol.
  12. Rinse twice with distilled water.
  13. Mount in glycerine water or glycerine jelly.

Expected Results

With appropriate filters, amyloid fluoresces bright yellow.

Notes

  • Although the method specifies an aqueous mounting medium, either blotting and treating with xylene repeatedly until clear, or dehydrating with ethanol and clearing with xylene, then mounting with a fluorescence free resinous medium may be satisfactory.

Safety Note

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

References

  1. Guntern, R., Bouras, C., Hof, P.R. & Vallett, P.G., (1992),
    An Improved Thioflavine S Method For Staining Neurofibrillary Tangles And Senile Plaques In Alzheimer’s Disease.
March 30, 2023
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Vassar & Culling’s Thioflavine T for Amyloid Fluorescence

By Amyloid, Fluorescent Staining, Protocols, Stain Target, Stain Type
Protocol

Vassar & Culling's Thioflavine T

for Amyloid Fluorescence

8
steps
3
materials
print

Materials

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 alum hematoxylin for 2 minutes.
  3. Rinse well with water.
  4. Place into thioflavine T solution for 3 minutes.
  5. Rinse with water.
  6. Place into acetic acid solution for 20 minutes.
  7. Wash with water.
  8. Mount in a fluorescence free aqueous 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 bright yellow.

Notes

  • The pretreatment with alum hematoxylin suppresses nuclear fluorescence.
  • Some workers have reported that materials other than amyloid may fluoresce yellow. The authors say this is caused by using a yellow barrier filter and strongly recommended the first filter combination. With this, these materials fluoresce white or pale blue.

Safety Note

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

References

  1. Vassar, P.S. and Culling, C.F.A., (1959),
    Fluorescent stains with special reference to amyloid and connective tissue,
    Archives of pathology, v 68, page 487
  2. 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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Birch-Hirschfeld’s Crystal Violet for Amyloid

By Amyloid, Metachromasia, Protocols, Stain Target, Stain Type
Protocol

Birch-Hirschfeld's Crystal Violet

for Amyloid

9
steps
4
materials
print

Materials

Tissue Sample

Frozen sections are preferred. Cryostat sections usually show brighter metachromasia. Unmounted frozen sections may also be floated in each solution and mounted on a slide just before coverslipping. 5µ paraffin sections of neutral buffered formalin fixed tissue are likely also suitable. Other fixatives may be satisfactory.

Protocol

  1. Bring sections to water via xylene and ethanol, except for cryostat and frozen sections.
  2. Place into bismarck brown solution for 5 minutes.
  3. Rinse well with 95% ethanol, then rinse with distilled water.
  4. Place into crystal violet solution for 5 minutes.
  5. Rinse with water.
  6. If necessary, differentiate in 1% acetic acid until amyloid is red and contrasts well with the tissue.
  7. Wash well in tap water.
  8. Drain all water from the slide until just damp and mount with levulose syrup.
  9. Ring the coverslip to inhibit evaporation of the mounting medium.

Expected Results

  • Amyloid  –  purple-red
  • Background  –  blue-violet
  • Nuclei  –  brown

Notes

  • Methyl violet may be used instead of crystal violet if preferred.
  • Although levulose syrup (fructose syrup or high fructose corn syrup) is specified it is likely that Highman’s gum syrup would be preferable as it inhibits leaching of the dye.
  • Although bismarck brown is metachromatic (yellow metachromasia), it is used here as a basic dye for staining nuclei.

Safety Note

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

References

  1. Gray, Peter. (1954)
    The Microtomist’s Formulary and Guide, p.451.
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
March 30, 2023
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Jurgens’ Crystal Violet for Amyloid

By Amyloid, Metachromasia, Protocols, Stain Target, Stain Type
Protocol

Jurgens' Crystal Violet

for Amyloid

7
steps
3
materials
print

Materials

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory. Cryostat sections usually show brighter metachromasia. Unmounted frozen sections may also be floated in each solution and mounted on a slide just before coverslipping.

Protocol

  1. Bring sections to water via xylene and ethanol, except for cryostat and frozen sections.
  2. Place into crystal violet solution for 2-5 minutes.
  3. Rinse well with water and examine microscopically.
  4. If necessary, differentiate in dilute acetic acid until amyloid is red and contrasts well with the tissue.
  5. Wash very well in tap water, about 5 minutes.
  6. Drain all water from the slide until just damp and mount with Highman’s medium.
  7. Ring the coverslip to inhibit evaporation of the mounting medium and precipitation of the ingredients.

Expected Results

  • Amyloid  –  purple-red
  • Background  –  blue-violet
  • Nuclei  –  blue-violet

Notes

  • Methyl violet may be used instead of crystal violet if preferred.
  • Highman’s gum syrup is a modification of Apathy’s gum syrup and contains potassium acetate or sodium chloride to stop bleeding of the dye into the mounting medium.

Safety Note

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

References

  1. Drury, R A, and Wallington, E A, (1967).
    Carleton’s histological technique., Ed. 4, p. 222.
    Oxford University Press, London, England.
March 30, 2023
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Anderson’s Alum Hematoxylin

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

Anderson's Alum Hematoxylin

8
steps
7
materials
print

Although there are three variations listed for Anderson’s formulas, those marked as 1923a and 1923b are variations of the same solution, the former being that given in the Microtomist’s Formulary and Guide and the latter in the Microtomist’s Vade Mecum.

Materials

MaterialVariantFunction
1923a1923b1929
Hematoxylin2.5 g2.5 g5 gDye
Ammonium alum20 gSat.30 gMordant
Distilled water900 mL900 mL700 mLSolvent
95% ethanol50 mL50 mL50 mLSolvent
Calcium hypochlorite4 g–40 gOxidant
Chloramine T or lime chloride4 gOxidant
Glacial acetic acid50 mL50 mL50 mLAcidifier

Compounding procedures

1923a & 1929

  1. Dissolve the calcium hypochlorite in 200 mL water.
  2. Dissolve the hematoxylin in some of the water.
  3. After 4 hours, combine the solutions.
  4. Dissolve the other ingredients in the rest of the water.
  5. Combine with the hematoxylin-hypochlorite solution.

1923b

  1. Bring 700 mL water to a boil, then saturate it with alum (see notes).
  2. Allow to cool for one day, then filter.
  3. Dissolve the chloramine T or lime chloride into 200 mL of water.
  4. Leave four hours. Shake occasionally.
  5. Dissolve the hematoxylin in the ethanol
  6. Add the oxidant solution to the hematoxylin.
  7. Mix for a few seconds. It should be dark brown.
  8. Slowly add to the 700 mL alum solution, while shaking.
  9. Add the acetic acid.
  10. It is ready immediately.

Protocol

  1. Bring sections to water with xylene and ethanol.
  2. Place into the staining solution for 2-3 minutes.
  3. Rinse well with water.
  4. Differentiate with acid ethanol if necessary.
  5. Rinse with water and blue.
  6. Rinse well with water.
  7. Counterstain if desired.
  8. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  blue
  • Background  –  as counterstain or unstained

Notes

  • The 1923a formula oxidizes 2.5 grams hematoxylin with 4 grams calcium hypochlorite. This is a ratio of 1.6:1.
  • The 1929 formula oxidizes 5 grams hematoxylin with 40 grams calcium hypochlorite. This is a ratio of 8:1.
  • The 1923b formula is an alternate formula given by Bolles-Lee. It differs from the 1923a formula by using a saturated alum solution, and chloramine T or “commercial chloride of lime” (a crude preparation of calcium chloride with other substances present) as oxidant.
  • The 1923b formula calls for 700 mL saturated aqueous alum. The instructions specify that the water should be saturated at boiling, then cooled to room temperature. This would require about 350 grams alum (at 500 mg/mL), but at room temperature, the solution would contain only about 100 grams (at 150 mg/mL). Dissolving 110 grams alum in 700 mL hot water, cooling and filtering would give the same solution.
  • Acid ethanol is 0.5% – 1% hydrochloric acid in 70% ethanol.
  • Blueing is done with alkaline solutions such as hard tap water, Scott’s tap water substitute, 0.1% ammonia water, 1% aqueous sodium acetate, 0.5% aqueous lithium carbonate etc.

Safety Note

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

References

  1. Gray, Peter. (1954)
    The Microtomist’s Formulary and Guide.
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
  2. Bolles Lee, A.. Edited by Gatenby, J.B. and Beams, H.W., (1950)
    The Microtomist’s Vade-Mecum. 11 ed.,
    Churchill, London, UK.
    Citing:
    Anderson, J., (1923)
    Journal of Pathology and Bacteriology.
  3. Susan Budavari, Editor, (1996)
    The Merck Index, Ed. 12
    Merck & Co., Inc., Whitehouse Station, NJ, USA
March 30, 2023
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Anderson’s Iron Hematoxylin

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

Anderson's Iron Hematoxylin

6
steps
7
materials
print

Materials

Solution A

MaterialAmountFunction
Hematoxylin0.5 gDye
100% ethanol50 mLSolvent
Distilled water50 mLSolvent
Calcium hypochlorite, 2% aqueous5 mLOxidant

Solution B

MaterialAmountFunction
Ferric ammonium sulphate3 gMordant
Distilled water100 mLSolvent
Sulfuric acid0.5 mLAcidifier

Compounding procedures

  1. Make each solution separately.
  2. For use, add 2 volumes of solution A to 1 volume of solution B.
  3. The working solution may be used immediately, but is not stable for long.

Protocol

  1. Bring sections to water with xylene and ethanol.
  2. Place into the staining solution for an appropriate time.
  3. Wash well in running tap water to blue.
  4. Rinse with distilled water.
  5. Counterstain if desired.
  6. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  black
  • Background  –  as counterstain or unstained

Notes

  • The stock solutions are stable for some time.
  • The working solution should be made fresh.
  • It is claimed that the working solution rarely overstains, i.e. it is progressive.
  • Solution A incorporates calcium hypochlorite as an oxidizing agent for hematoxylin. Presumably other oxidizing agents would suffice equally as well. Sodium iodate (0.1 g or less) is the most common.
  • The staining time should be determined by trial.

Safety Note

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

References

  1. Gray, Peter. (1954)
    The Microtomist’s Formulary and Guide.
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
    Citing:
    Anderson, J., (1929)
    How to stain the nervous system.
    Livingstone. Edinburgh, Scotland.
March 30, 2023
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Apathy’s Alum Hematoxylin

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

Apathy's Alum Hematoxylin

8
steps
7
materials
print

Materials

MaterialAmountFunction
Hematoxylin3 gDye
Ammonium alum30 gMordant
Distilled water450 mLSolvent
95% ethanol250 mLDye solvent
Glycerol350 mLStabiliser
Glacial acetic acid10 mLAcidifier
Salicylic acid0.3 gAcidifier

Compounding procedure

  1. Dissolve the dye in 100 mL water and 250 mL ethanol.
  2. Leave at room temperature to ripen (months).
  3. Dissolve the alum and acids in 350 mL water.
  4. Combine both solutions.
  5. Add glycerol.

Protocol

  1. Bring sections to water with xylene and ethanol.
  2. Place into the staining solution for an appropriate time.
  3. Rinse well with water.
  4. Differentiate with acid ethanol if necessary.
  5. Rinse with water and blue.
  6. Rinse well with water.
  7. Counterstain if desired.
  8. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  blue
  • Background  –  as counterstain or unstained

Notes

  • Lillie excludes the salicylic acid.
  • The appropriate staining time should be determined by trial.
  • Acid ethanol is 0.5% – 1% hydrochloric acid in 70% ethanol.
  • Blueing is done with alkaline solutions such as hard tap water, Scott’s tap water substitute, 0.1% ammonia water, 1% aqueous sodium acetate, 0.5% aqueous lithium carbonate etc.

Safety Note

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

References

  1. Gray, Peter. (1954)
    The Microtomist’s Formulary and Guide.
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
  2. Lillie, R.D., (1954)
    Histopathologic technique and practical histochemistry Ed.2
    Blakiston, New York, USA.
March 30, 2023
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