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Stain Target

Highman’s Crystal Violet for Amyloid

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

Highman's Crystal Violet

for Amyloid

7
steps
5
materials

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. Stain nuclei with Weigert’s iron hematoxylin for 5 minutes.
  3. Wash with water.
  4. Place into crystal violet solution for 1-30 minutes until amyloid is stained.
  5. Rinse well with water.
  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 – black

Notes

  • Methyl violet may be used instead of crystal violet if preferred.
  • Gray notes that Lieb substituted a solution of 0.3% crystal violet in 0.3% hydrochloric acid for Highman’s crystal violet solution.
  • 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. Gray, Peter. (1954)
    The Microtomist’s Formulary and Guide, p. 452.
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.

Lendrum’s’ Methyl Violet for Amyloid

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

Lendrum's' Methyl Violet

for Amyloid

7
steps
3
materials

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 methyl violet solution for 3 minutes.
  3. Differentiate in formalin until amyloid is red and contrasts well with the tissue.
  4. Place into sodium chloride solution for 5 minutes.
  5. Rinse well with tap water.
  6. Drain all water from the slide until just damp and mount with corn syrup.
  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

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), p. 466.
    Cellular pathology technique. Ed. 4.,
    Butterworths, London, England.

King’s Silver Impregnation for Amyloid

By Amyloid, Metal Impregnation, Metal Impregnation, Silver, Protocols, Stain Target, Stain Type
Protocol

King's Silver Impregnation

for Amyloid

7
steps
10
materials

Materials

  • Pyridine
  • Ethanol 50%, 75%, 95%
  • Sodium carbonate
    MaterialAmount
    Sodium carbonate, anhydrous3.5g
    Distilled water100mL
  • Silver nitrate, 10%
    MaterialAmount
    Silver nitrate10g
    Distilled water100mL
  • Carbol xylene
    MaterialAmount
    Xylene3volumes
    Phenol1volume
  • Ammoniacal Silver
    MaterialAmount
    Silver nitrate, 10% aqueous5mL
    Strong ammonia (S.G. 0.880)asrequired
    Sodium carbonate, aqueous6.8mL
    Distilled waterasrequired

    Add ammonium hydroxide drop by drop to 5 mL of the silver solution until the precipitate which forms just redissolves. Add 6.8 mL sodium carbonate solution and sufficient water to make up to 75 mL. For use, add a few drops of pyridine to each 10 mL of the solution.

Tissue Sample

10-15µ free floating frozen sections of formalin fixed tissues. Collect into water and wash in a few changes of distilled water to remove residual formalin.


Protocol

  1. Add pyridine to 10 mL ammoniacal silver solution and warm to 40°C.
  2. Float sections in the warm ammoniacal silver until they turn brown.
  3. Rinse with water.
  4. Place in 50% ethanol for 5-10 minutes.
  5. Quickly treat with 75% and 95% ethanols.
  6. Clear with carbol xylene.
  7. Mount sections on to slides and coverslip, using a resinous medium.

Expected Results

  • Amyloid – brown to black.

Safety Note

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


References

  1. King, L.S., (1948),
    Atypical Amyloid Disease: With Observation on a New Silver Stain for Amyloid, American Journal of Pathology, v 24, page 1095-1115

Lynch & Inwood’s Gold for Amyloid

By Amyloid, Metal Impregnation, Metal Impregnation, Non-Silver, Protocols, Stain Target, Stain Type
Protocol

Lynch & Inwood's Gold

for Amyloid

7
steps
5
materials

Materials

  • Chloro-auric acid, 1% aqueous
  • Hydrogen peroxide, 3% aqueous
  • Iodine solution
    MaterialAmount
    Iodine1g
    Potassium iodide2g
    Distilled water100mL

    Mix the potassium iodide and iodine crystals together. Add 5 mL of the water and mix until both have dissolved. Add the rest of the water.

Tissue Sample

5-10µ sections from formalin fixed, paraffin embedded tissues, mounted on slides are suitable.


Protocol

  1. Bring sections to water through xylene and ethanols.
  2. Place in the iodine solution for 2½-5 minutes.
  3. Rinse with distilled water, 3 times of 5-10 seconds each.
  4. Place in chloro-auric acid solution for 2½-5 minutes.
  5. Rinse with distilled water, 3 times of 5-10 seconds each.
  6. Place in fresh 3% hydrogen peroxide at 37°C for 6-36 hours.
  7. Dehydrate with ethanol, clear with xylene and mount using a resinous medium.

Expected Results

  • Amyloid – golden yellow to faint purple.
  • Erythrocytes, muscle and liver – blue
  • Nuclei and cytoplasm – brown
  • Collagen – grey

Safety Note

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


References

  1. Lynch, M.J. and Inwood, M.J.H., (1963),
    Gold as a permanent stain for amyloid,
    Stain Technology, v 38, page 260.

Pappenheim Stain for Plasma Cells

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

Pappenheim Stain

for Plasma Cells

5
steps
4
materials

Materials

Stock solution A

MaterialAmount
Methyl green1g
Distilled water100mL
Phenol0.25g

Stock solution B

MaterialAmount
Pyronin Y4g
Distilled water100mL
Phenol5g

Working solution

MaterialAmount
Stock solution A15mL
Stock solution B35mL

Tissue Sample

Most fixatives should be satisfactory.


Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into the staining solution for 5-10 minutes.
  3. Rinse briefly with water.
  4. Dehydrate rapidly with acetone.
  5. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  violet
  • Plasma cell cytoplasm  –  red
  • Other cell cytoplasm  –  pink

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. 355
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.

Sandifords Stain for Plasma Cells

By Intracytoplasmic Granules, Plasma Cells, Protocols, Stain Target

Sandifords Stain

for Plasma Cells

5
steps
6
materials

Materials

Staining solution

MaterialAmount
Distilled water75mL
Glycerol20mL
Ethanol, 95%5mL
Methyl green0.15g
Pyronin Y0.5g
Phenol1.5g

Tissue Sample

Most fixatives should be satisfactory if fixation is not extended. Reagents which cause depolymerisation of DNA should be avoided. Decalcification may interfere with staining.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into the staining solution for 5-10 minutes.
  3. Rinse briefly with water.
  4. Dehydrate rapidly with acetone.
  5. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  violet
  • Plasma cell cytoplasm  –  red
  • Other cell cytoplasm  –  pink

Notes

  • The reference gave no details of the method to use. The details above are from that given for Pappenheim’s solution, and should be suitable as a starting point.
  • Time and temperature of staining may need to be increased.

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. 355
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.

Unna’s Stain for Plasma Cells

By Intracytoplasmic Granules, Plasma Cells, Protocols, Stain Target

Unna's Stain

for Plasma Cells

5
steps
6
materials

Materials

Staining Solution

MaterialAmount
Distilled water100mL
Phenol0.5g
Glycerol20mL
Ethanol, absolute2.5mL
Pyronin Y0.25g
Methyl green0.15g

Preparation

  1. Place both dyes in a mortar, add the ethanol and grind together.
  2. Heat the glycerol to 50°C and add small volumes to the dyes while grinding.
  3. Dissolve the phenol in the water and wash the dyes from the mortar. Filter.

Tissue Sample

Most fixatives should be satisfactory.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into the staining solution at 30°C for 10 minutes.
  3. Rinse briefly with water.
  4. Place in absolute ethanol until differentiated.
  5. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  violet
  • Plasma cell cytoplasm  –  red
  • Other cell cytoplasm  –  pink

Notes

  • In their description of this method, Carleton and Leach omit the method of preparation, implying that the trituration of the dyes may not be necessary.

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. 355
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
  2. Carleton, H M, and Leach, E H, (1938).
    Histological technique., Ed. 2.
    Oxford University Press, London, England.

Phenolic MGP Solutions

By Intracytoplasmic Granules, Plasma Cells, Protocols, Stain Target

Phenolic MGP Solutions

7
steps
7
materials

Materials

Table 1. Comparison of phenolic methyl green –pyronin solutions for plasma cells

MaterialFormula
PappenheimUnnaSandifordScott & FrenchLangeronKurnick
Distilled water100mL82mL75mL80mL100mL100mL
GlycerolmL16mL20mL16mLmLmL
Ethanol, 100%0mL2mL0mL4mLmLmL
Ethanol, 95%mLmL5mLmLmLmL
Phenol0.25g0.4g1.5g1.6g5gg
Methyl green0.3g0.12g0.15g0.8g2g0.6g
Pyronin Y0.7g0.2g0.5g0.2g2g1g

Tissue Sample

Most fixatives should be satisfactory.

Protocol

These solutions have been included for comparison purposes, and the formulae have been adjusted to make 100 mL of each solution to facilitate the comparison. See the individual methods for details of their use and the actual formula. There is no single method for their application, but the details below may be used as a starting point.

  1. Bring sections to water via xylene and ethanol.
  2. Place in the MGP solution for 10 minutes at room temperature.
  3. Rinse with distilled water.
  4. Differentiate with absolute ethanol if required.
  5. Complete dehydration with acetone if necessary.
  6. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  blue-green
  • Plasma cell cytoplasm  –  red
  • Other cell cytoplasm  –  pink

Notes

  • The time and temperature in step 2 may need to be increased.
  • Differentiation may not be required, depending on the time and temperature of staining.
  • Various fluids have been used for dehydration: acetone, n-butanol, t-butanol, 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. p. 355
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
  2. Disbrey, B. D., (1970)
    Histological laboratory methods.
    E. & S. Livingstone, Edinburgh and London, UK.

Robb-Smith’s Impregnation for Reticulin

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

Robb-Smith's Impregnation

for Reticulin

20
steps
10
materials

Materials

  • Silver nitrate, 5% aqu.
  • Silver nitrate, 10% aqu.
  • Sodium hydroxide, 10% aqu.
  • Strong ammonium hydroxide (s.g. 0.88)
  • Ammonium hydroxide 10% aqu.
  • Potassium permanganate, 0.25% aqu.
  • Oxalic acid, 1.5% aqu.
  • Formalin, 37.5% aqu.
  • Yellow gold chloride, 0.2% aqu.
  • Sodium thiosulphate, 5% aqu.

Preparation of Robb-Smith’s Ammoniacal Silver

  1. Place 28.5 mL of 10% aqueous silver nitrate in a flask.
  2. Add 0.6 mL of 10% aqueous sodium hydroxide
  3. Add strong ammonium hydroxide by drops until the precipitate just dissolves.
  4. The solution should have a faint opalescence.
  5. Dilute to 100 mL with distilled water.

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue mounted on slides are suitable. Other fixatives are likely to be satisfactory. A section adhesive is recommended. The paraffin wax does not need to be removed.

Protocol

  1. Place sections with the paraffin wax intact in 10% ammonia for 15 minutes
  2. Wash with tap water.
  3. Oxidize with 0.25% potassium permanganate for 5 minutes.
  4. Wash with tap water.
  5. Place in oxalic acid until bleached.
  6. Wash with tap water.
  7. Place in 5% silver nitrate for 1 hour.
  8. Rinse with distilled water.
  9. Treat with ammoniacal silver solution for 15 minutes.
  10. Wash with distilled water.
  11. Place in 37.5% formalin for 3 minutes.
  12. Wash with tap water.
  13. Rinse with distilled water.
  14. Tone with 0.2% gold chloride solution for 3 minutes.
  15. Rinse with distilled water.
  16. Fix in 5% sodium thiosulphate for 1 minute.
  17. Wash well with running tap water.
  18. Dehydrate with ethanol.
  19. Remove paraffin wax and clear with a few changes of xylene.
  20. Mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Background  –  grey

Notes

  • The 10% ammonium hydroxide is made by diluting 1 part strong ammonium hydroxide with 9 parts distilled water.
  • The 37.5% formalin solution (15% formaldehyde) is made by diluting 37.5 mL strong formalin with 62.5 mL tap water.
  • Ensure that the strong ammonium hydroxide is fresh and full strength. Keep well stoppered when not in use. Pour sufficient for use into a beaker, then immediately restopper the bottle. Do not return unused solution to the stock 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.
  • 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

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

Studricka’s Impregnation for Reticulin in Decalcified Tissue

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

Studricka's Impregnation

for Reticulin in Decalcified Tissue

12
steps
7
materials

Materials

  • Silver nitrate, 10% aqu.
  • Silver nitrate, 3% aqu.
  • Strong ammonium hydroxide (s.g. 0.88)
  • Sodium hydroxide, 10% aqu.
  • Formalin, 10% aqu.
  • Yellow gold chloride, 0.5% aqu.
  • Sodium thiosulphate, 5% aqu.

Preparation of Studricka’s Ammoniacal Silver

  1. Place 100 mL of 10% aqueous silver nitrate in a flask.
  2. Add 10% aqueous sodium hydroxide until no more precipitate forms (with a Pasteur pipette).
  3. Add strong ammonium hydroxide by drops until the precipitate just dissolves.

Tissue Sample

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

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place in 3% silver nitrate for 4 days.
  3. Wash with distilled water.
  4. Place in ammoniacal silver solution until sections are yellow-brown.
  5. Rinse with distilled water.
  6. Place in 10 formalin until sections are dark brown.
  7. Wash with distilled water.
  8. Tone with 0.5% gold chloride until sections are grey-black.
  9. Wash with distilled water.
  10. Fix in 5% sodium thiosulphate for 1 hour.
  11. Wash well with running tap water.
  12. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Background  –  grey

Notes

  • Ensure that the strong ammonium hydroxide is fresh and full strength. Keep well stoppered when not in use. Pour sufficient for use into a beaker, then immediately restopper the bottle. Do not return unused solution to the stock 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.
  • 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.
  • Gray notes that Zimmerman recommended treatment with 3% silver nitrate for 2 days (step 2), and that the ammoniacal silver solution should be diluted 1 part with 3 parts distilled water (step 4).

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.