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Buffered Thionin for Nissl Bodies

By Intracytoplasmic Granules, Nissl Bodies, Protocols, Stain Target
Protocol

Buffered Thionin

for Nissl Bodies

16
steps
3
materials
print

Materials

MaterialAmount for pH 3.7 SolutionAmount for pH 4.5 Solution
Acetic acid, 0.6% (0.1M)90mL60mL
Sodium acetate, 0.8% (0.1M)10mL40mL
Thionin, 1% aqueous2.5mL2.5mL

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

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

Alternative Method

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

Expected Results

StructurepH 3.7 Staining SolutionpH 4.5 Staining Solution
Nissl bodiesbluedark blue
Nucleibluedark blue
Backgroundpale or unstainedpale blue

Safety Note

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


References

  1. 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 Amyloid, Fluorescent Staining, Protocols, Stain Target, Stain Type
Protocol

Burns, Pennock & Stoward's Thioflavine T

for Amyloid Fluorescence

6
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. Stain nuclei with alum hematoxylin.
  3. Rinse well with water.
  4. Stain with thioflavine T solution for 5 minutes.
  5. Blot, rinse well with absolute ethanol.
  6. 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

  1. 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.
  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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Bullard’s Alum Hematoxylin

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

Bullard's Alum Hematoxylin

8
steps
10
materials
print

Materials

MaterialAmountFunction
Hematoxylin8 gDye
Ammonium alum20 gMordant
Distilled water250 mLSolvent
50% ethanol144 mLSolvent
Glacial acetic acid16 mLAcidifier
Mercuric oxide, red8 gOxidant
95% ethanol275 mLSolvent
Glycerol330 mLStabiliser
Glacial acetic acid18 mLAcidifier
Ammonium alum40 gMordant

Compounding procedure

  1. Combine the first five ingredients and bring to a boil.
  2. Add the mercuric oxide with caution.
  3. Cool and filter.
  4. Add the last four ingredients.
  5. The solution may be used immediately, and is stable for more than a year.

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

  • The solution is likely regressive
  • The appropriate 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. F. A. Putt
    Manual of Histopathological Staining Methods
    John Wiley & Sons, New York, NY., USA
March 30, 2023
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Roque’s Trichrome for Mallory Bodies

By Protocols, Stain Type, Trichrome Staining, Trichrome, One-Step
Protocol

Roque's Trichrome

for Mallory Bodies

9
steps
6
materials
print

Materials

  • Weigert’s iron hematoxylin or equivalent
  • Solution A
    MaterialAmount
    Phosphomolybdic acid1g
    Distilled water100mL
  • Solution B
    MaterialAmount
    Aniline blue0.5g
    Chromotrope 2R2g
    Hydrochloric acid, 0.02N100mL

    Dissolve the aniline blue in the acid with gentle heat. Cool. Add the chromotrope 2R. Filter.

Tissue Sample

3µ paraffin sections of neutral buffered formalin fixed liver 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

  1. Bring sections to water via xylene and ethanol.
  2. Stain nuclei with Weigert’s iron hematoxylin or equivalent for 5 minutes.
  3. Wash in tap water to blue.
  4. Place into solution A for 2 minutes.
  5. Rinse with distilled water.
  6. Place into solution B for 8 minutes.
  7. Rinse with distilled water.
  8. Dehydrate with ethanol.
  9. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  black
  • Cytoplasm  –  red
  • Muscle  –  red
  • Mallory bodies  –  bright blue, or red with blue periphery
  • Collagen  –  blue

Safety Note

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


References

  1. Roque, Agustin L., (1953),
    Chromotrope aniline blue method of staining Mallory bodies of Laennec’s cirrhosis.
    Laboratory Investigation, v. 2, no. 5, pp. 15-21.
March 30, 2023
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Carazzi’s Alum Hematoxylin

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

Carazzi's Alum Hematoxylin

6
steps
5
materials
print

Materials

MaterialAmountFunction
Hematoxylin1 gDye
Potassium alum50 gMordant
Distilled water800 mLSolvent
Glycerol200 mLStabiliser
Sodium iodate0.2 gOxidant

Compounding procedure

  1. Dissolve the hematoxylin in the glycerol.
  2. Dissolve the alum in 750 mL of the water.
  3. Dissolve the sodium iodate in the remaining 50 mL water.
  4. Add the alum solution to the hematoxylin solution slowly, while mixing well.
  5. Add the sodium iodate solution. Mix well.
  6. Filter.
  7. The solution may be used immediately, and is stable for about six months.

Protocol

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

Expected Results

  • Nuclei  –  blue
  • Background  –  as counterstain or unstained

Notes

  • This is a progressive solution giving little background staining.
  • Doubling the hematoxylin to 2 g intensifies nuclear staining.
  • The double strength solution is recommended for frozen sections with about 1 minute staining time.
  • 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.
  • In the original paper, 0.02 grams sodium iodate was specified. This would oxidize only a small part of the hematoxylin. The amount specified above would permit full oxidation.

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.
  2. Gray, Peter. (1954)
    The Microtomist’s Formulary and Guide. p.187.
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
March 30, 2023
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Llewellyn’s Sirius Red for Amyloid

By Amyloid, Direct Dye Staining, Eosinophils, Intracytoplasmic Granules, Paneth Cells, Protocols, Stain Target, Stain Type
Protocol

Llewellyn's Sirius Red

for Amyloid

8
steps
3
materials
print

Materials

MaterialAmount
Sirius red F3B0.5g
Distilled water50mL
Ethanol, absolute50mL

Staining Solution Preparation

  1. Dissolve the dye into the water, add ethanol and mix well.
  2. 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

  1. Bring sections to water via xylene and ethanol.
  2. Stain nuclei with a progressive alum hematoxylin for a few minutes.
  3. Rinse with tap water.
  4. Rinse with ethanol.
  5. Place into alkaline sirius red for 1 – 2 hours.
  6. Rinse well with tap water.
  7. Dehydrate with absolute ethanol.
  8. 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

  1. 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 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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