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Stain Target

Wilder’s Impregnation for Reticulin

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

Wilder's Impregnation

for Reticulin

19
steps
12
materials
print

Materials

  • Silver nitrate, 2% aqu.
  • Strong ammonium hydroxide (s.g. 0.88)
  • Sodium hydroxide, 40% aqu.
  • Potassium permanganate, 1% aqu.
  • Oxalic acid, 5% aqu.
  • Uranium nitrate, 1% aqu.
  • Yellow gold chloride, 0.2% aqu.
  • Sodium thiosulphate, 5% aqu.
  • Neutral red, 1% aqu.
  • Developer
    MaterialAmount
    Strong formalin5mL
    Uranium nitrate0.15g
    Water1L

Preparation of Bielchowsky’s Ammoniacal Silver

  1. Place 48 mL of 2% silver nitrate in a flask.
  2. Add 0.4 mL of 40% sodium hydroxide.
  3. While swirling, slowly add drops of strong ammonium hydroxide until the precipitate just redissolves.
  4. Make up to 100 mL with distilled water.

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory, but see the notes for comments about Zenker and Helly type fixed tissue. A section adhesive is recommended.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Oxidize with 1% potassium permanganate for 1 minute.
  3. Rinse well with tap water.
  4. Bleach in Oxalic acid for 1 minute.
  5. Rinse well with tap water.
  6. Sensitize with 1% uranium nitrate solution for 5-10 seconds.
  7. Rinse with distilled water.
  8. Treat with Bielchowsky’s ammoniacal silver for 1 minute.
  9. Rinse briefly with 90% ethanol.
  10. Place in the developer for 1 minute.
  11. Rinse well with tap water.
  12. Rinse with distilled water.
  13. Tone with 0.2% gold chloride solution.
  14. Rinse with distilled water.
  15. Fix in 5% sodium thiosulphate for 1 minute.
  16. Wash well with running tap water.
  17. Counterstain with neutral red for 1 minute.
  18. Rinse with tap water.
  19. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Nuclei  –  red
  • Background  –  grey

Notes

  • For Zenker and Helly type fixatives, Wilder recommended a slightly different procedure.
    • Replace the Mallory bleach (steps 2-4) with 10% phosphomolybdic acid treatment for 1 minute, followed by a water rinse.
    • Use Foot’s ammoniacal silver solution instead of Bielchowsky’s.

      Foot’s ammoniacal silver: To 10 mL of 1% silver nitrate in a flask, add 0.1 mL of 40% potassium hydroxide. Add strong ammonium hydroxide drop by drop while shaking the solution until the precipitate just dissolves. Make up to 100 mL with distilled water.

  • Ensure that both the ammonium hydroxide and sodium hydroxide are fresh and full strength. Keep both well stoppered when not in use. For the ammonium hydroxide, pour sufficient for use from the stock bottle into a beaker, then immediately restopper the stock bottle. Do not return excess ammonium hydroxide to the stock bottle.
  • After making the ammoniacal silver solution, smell the solution to ensure it has only a faint smell of ammonia. If the smell of ammonia is strong, it indicates that too much ammonium hydroxide has been added. If so, it is preferable to make the solution again. Improperly made ammoniacal silver solutions can affect the quality of the impregnation.
  • It is sometimes difficult to obtain uranium nitrate, particularly if it requires international transportation.
  • The strong formalin used to make the developer should be neutralized, but do not use buffered formalin. Neutral formalin in this context may be made by keeping strong formalin over marble chips. However, be very careful as the gas given off may increase the pressure inside the container and cause an explosion. Either apply a cap loosely so gas can escape, or use a fermentation lock.
  • 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.
March 30, 2023
Love0

Zimmerman’s Impregnation for Reticulin in Decalcified Tissue

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

Zimmerman's Impregnation

for Reticulin in Decalcified Tissue

12
steps
7
materials
print

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 Zimmerman’s Ammoniacal Silver

  1. Place 25 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.
  4. Dilute to 100 mL with distilled water.

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 2 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.
  • This method is a modification of Studricka’s technique using a shorter time in silver nitrate (step 2), and a more dilute ammoniacal silver solution (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.
March 30, 2023
Love0

Maresch’ Impregnation for Reticulin in Liver

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

Maresch' Impregnation

for Reticulin in Liver

12
steps
8
materials
print

Materials

  • Silver nitrate, 2% aqu.
  • Silver nitrate, 10% aqu.
  • Sodium hydroxide, 40% aqu.
  • Strong ammonium hydroxide (s.g. 0.88).
  • Formalin, 20% aqu.
  • Yellow gold chloride, 0.2% in 0.2% acetic acid.
  • Sodium thiosulphate, 5% aqu.
  • Neutral red, 1% aqu.

Preparation of Maresch’ Ammoniacal Silver

  1. Place 100 mL of 10% silver nitrate in a flask.
  2. Add 0.5 mL of 40% sodium hydroxide and mix well.
  3. Add drops of strong ammonium hydroxide until the precipitate just dissolves.

Tissue Sample

5 µ paraffin sections of neutral buffered formalin fixed 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 2% silver nitrate for 12-24 hours.
  3. Treat with ammoniacal silver solution for 2-30 minutes.
  4. Rinse well with distilled water.
  5. Place in 10% formalin for 1 hour.
  6. Wash well with tap water.
  7. Rinse with distilled water.
  8. Tone with acidified gold chloride solution for 1 hour.
  9. Rinse well with distilled water.
  10. Fix in 5% sodium thiosulphate for 10-15 minutes.
  11. Wash well with tap water.
  12. Dehydrate with ethanol, clear with carbol-xylene, and mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Background  –  grey

Notes

  • 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.
  • 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.
March 30, 2023
Love0

Nassar & Shanklin Impregnation for Reticulin

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

Nassar & Shanklin Impregnation

for Reticulin

22
steps
12
materials
print

Materials

  • Silver nitrate, 10% aqu.
  • Strong ammonium hydroxide (s.g. 0.880).
  • Oxalic acid, 2% aqu.
  • Yellow gold chloride, 2% aqu.
  • Sodium thiosulphate, 5% aqu.
  • Progressive hemalum
  • Mallory bleach
    MaterialAmount
    Potassium permanganate, 1% aqu.1vol
    Sulphuric acid, 0.5% aqu.1vol
  • Pyridinised silver nitrate
    MaterialAmount
    Silver nitrate, 2% aqu.10mL
    Pyridine3 drops
  • Reducer
    MaterialAmount
    Formalin, 2% aqu. neutralised1vol
    Ethanol, 95%1vol

Preparation of Ammoniacal Silver

  1. Place 1 mL of strong ammonium hydroxide into a flask.
  2. Pour in 7 mL of 10% aqueous silver nitrate.
  3. Add drops of 10% silver nitrate until a persistent, pale opalescence remains after shaking.
  4. Add an equal volume of distilled water. Add 3 drops pyridine for each 10 mL of solution for use.

Tissue Sample

5 µ paraffin sections of neutral buffered formalin fixed 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. Oxidise with the Mallory bleach for 1-2 minutes.
  3. Rinse with distilled water.
  4. Bleach in Oxalic acid for 1-2 minutes.
  5. Rinse with distilled water.
  6. Rinse with tap water for 5 minutes.
  7. Rinse with 95% ethanol.
  8. Place in 2% silver nitrate
  9. Rinse with 95% ethanol.
  10. Treat with pyridinised silver nitrate at 50°C for 30 to 60 minutes.
  11. Rinse briefly with 95% ethanol.
  12. Treat with ammoniacal silver at 50°C for 5 minutes.
  13. Rinse briefly with 95% ethanol.
  14. Treat with the reducer for 2 minutes.
  15. Rinse well with distilled water.
  16. Tone with gold chloride solution until sections are grey.
  17. Rinse well with distilled water.
  18. Fix in 5% sodium thiosulphate for 2 minutes.
  19. Wash well with running tap water.
  20. Counterstain with a progressive hemalum and blue.
  21. Wash with tap water.
  22. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Nuclei  –  blue
  • Background  –  grey

Notes

  • Ensure that the ammonium hydroxide is fresh and full strength. Keep well stoppered when not in use. Remove sufficient for use from the stock bottle then immediately restopper.
  • 2% formalin is made by diluting strong formalin 1:50 with tap water (2 mL strong formalin, 98 mL tap water). The strong formalin used to make this should be neutralised, but do not use buffered formalin. Neutral formalin in this context may be made by keeping strong formalin over marble chips. However, be very careful as the gas given off may increase the pressure inside the container and cause an explosion. Either apply a cap loosely so gas can escape, or use a fermentation lock.
  • 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. Drury, R A, and Wallington, E A, (1967).
    Carleton’s histological technique., Ed. 5.
    Oxford University Press, London, England.
March 30, 2023
Love0

Nissl’s Methylene Blue for Nissl bodies

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

Nissl's Methylene Blue

for Nissl bodies

7
steps
5
materials
print

Materials

  • Cajeput oil
  • Staining solution
    MaterialAmount
    Castile soap1.75g
    Methylene blue3.75g
    Distilled water1L

    Dissolve the soap in the water. Add and dissolve the dye. Allow to ripen at least three months.

  • Differentiator
    MaterialAmount
    Aniline10mL
    Ethanol, 95%90mL

Tissue Sample

Ethanol fixation is preferred. Formalin fixed tissue may be suitable. Sections should be thicker than usual, and were originally to be free floating sections, likely free hand sections prepared without freezing.

Protocol

  1. Place sections in the staining solution in a watch glass.
  2. Heat gently until bubbles appear.
  3. Transfer to the differentiating fluid in another watch glass.
  4. Differentiate until color ceases to be extracted.
  5. Transfer the section to a slide and gently blot dry.
  6. Clear with cajeput oil.
  7. Mount with a resinous medium.

Expected Results

  • Nissl  –  blue
  • Nuclei  –  blue

Notes

  • Nissl specified “Venetian soap”, which is the same as “Castile soap”. It refers to solid, bar soap made from olive oil and sodium hydroxide.

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. pp. 446.
    Originally published by: The Blakiston Co.
    Republished by: Robert E. Krieger Publishing Co.
  2. Gatenby, J.B. and Beams, H.W., (1950)
    The Microtomist’s Vade-Mecum. 11 ed., pp. 508, para. 1099
    Churchill, London, UK.
March 30, 2023
Love0

Toluidine Blue for Nissl bodies

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

Toluidine Blue

for Nissl bodies

7
steps
7
materials
print

Materials

  • Staining Solution
    • Option 1
      MaterialAmount
      Toluidine blue1g
      Sodium tetraborate1g
      Distilled water100mL
    • Option 2
      MaterialAmount
      Thionin0.1g
      Distilled water100mL
  • Gothard’s Differentiator
    MaterialAmount
    Creosote50mL
    Cajeput oil40mL
    Xylene50mL
    Ethanol, absolute160mL

Tissue Sample

10µ 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 into the staining solution at 56°C for at least 30 minutes
    or up to overnight at room temperature.
  3. Rinse well with running tap water.
  4. Rinse with absolute ethanol.
  5. Differentiate with Gothard’s differentiator, controlling microscopically.
  6. Rinse well with absolute ethanol.
  7. Clear with xylene and mount using a resinous medium.

Expected Results

  • Nissl bodies  –  blue
  • Nuclei  –  blue
  • Background  –  pale to unstained

Notes

  • Disbrey omits the sodium tetraborate from the toluidine blue solution and stains at room temperature, but recommends overnight staining when possible.
  • Methylene blue may be substituted for thionin.
  • Culling recommends diluting the Gothard’s differentiator with an equal volume of absolute ethanol before use.

Safety Note

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

References

  1. Disbrey, B. D., (1970)
    Histological laboratory methods, p. 232.
    E. & S. Livingstone, Edinburgh and London, UK.
  2. Culling, C F A, Allison, R T, Barr, W T, (1985).
    Cellular pathology technique., Ed. 4.
    Butterworths, London, England.
March 30, 2023
Love0

Laidlaw’s Impregnation for Reticulin on Bouin Fixed Tissue

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

Laidlaw's Impregnation

for Reticulin on Bouin Fixed Tissue

20
steps
10
materials
print

Materials

  • Iodine, 1% in 95% ethanol
  • Sodium thiosulphate, 5% aqu.
  • Potassium permanganate, 0.5% aqu.
  • Oxalic acid, 5% aqu.
  • Formalin, 1% aqu.
  • Yellow gold chloride, 0.2% aqu.
  • Silver nitrate, 60% aqu.
  • Strong ammonium hydroxide (s.g. 0.88)
  • Lithium carbonate, saturated aqu.
  • Neutral red, 1% aqu.

Preparation of Ammoniacal Silver

  1. Place 16 mL of the 60% silver nitrate in a flask.
  2. Add 185 mL of saturated lithium carbonate.
  3. Allow the precipitate to settle and decant the supernatent.
  4. Wash the precipitate with water, allow to settle, and decant several times.
  5. Add 60 mL distilled water.
  6. Add drops of strong ammonium hydroxide until the precipitate just redissolves.
  7. Dilute to 100 mL with distilled water.

Tissue Sample

5 µ paraffin sections of Bouin’s fluid fixed tissue are suitable. Other fixatives are likely to be satisfactory. A section adhesive is recommended. The method was intended for staining reticulin in skin, but there is no reason it would not be satisfactory for other tissues.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place in alcoholic iodine for 3 minutes.
  3. Rinse well with tap water.
  4. Bleach with sodium thiosulphate for 3 minutes.
  5. Oxidise with potassium permanganate for 3 minutes.
  6. Rinse well with tap water.
  7. Bleach in Oxalic acid for a 5 minutes.
  8. Rinse well with tap water.
  9. Rinse with distilled water.
  10. Treat with ammoniacal silver for 10 minutess.
  11. Rinse with distilled water.
  12. Reduce in formalin for 10 minute.
  13. Rinse well with tap water.
  14. Rinse with distilled water.
  15. Tone with 0.2% gold chloride for 10 minutes.
  16. Rinse with distilled water.
  17. Wash well with running tap water.
  18. Counterstain with neutral red for 1 minute.
  19. Rinse with tap water.
  20. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Nuclei  –  red
  • Background  –  grey

Notes

  • Note that this method does not say to fix the impregnation with sodium thiosulphate. If sections fade or fixing the impregnation is considered desirable, then Place in 5% sodium thiosulphate for 5 minutes after step 16.
  • Ensure that both the ammonium hydroxide and sodium hydroxide are fresh and full strength. Keep both well stoppered when not in use. For the ammonium hydroxide, pour sufficient for use from the stock bottle into a beaker, then immediately restopper the stock bottle. Do not return excess ammonium hydroxide to the stock bottle.
  • After making the ammoniacal silver solution, smell the solution to ensure it has only a faint smell of ammonia. If the smell of ammonia is strong it indicates that too much ammonium hydroxide has been added. If so, it is preferable to make the solution again. Improperly made ammoniacal silver solutions can affect the quality of the impregnation.
  • 1% formalin is made by diluting strong formalin 1:100 with tap water (1 mL strong formalin, 99 mL tap water).
  • 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.
March 30, 2023
Love0

Levi’s Impregnation for Reticulin in Zenker fixed Lymph Nodes

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

Levi's Impregnation

for Reticulin in Zenker fixed Lymph Nodes

14
steps
8
materials
print

Materials

  • Silver nitrate, 2% aqu.
  • Silver nitrate, 20% aqu.
  • Sodium hydroxide, 40% aqu.
  • Strong ammonium hydroxide (s.g. 0.88).
  • Formalin, 12% aqu.
  • Yellow gold chloride, 0.5% aqu.
  • Sodium thiosulphate, 5% aqu.
  • Neutral red, 1% aqu.

Preparation of Levi’s Ammoniacal Silver

  1. Place 17 mL of 20% silver nitrate in a flask.
  2. Add 17 mL of 40% sodium hydroxide and mix well.
  3. Add drops of strong ammonium hydroxide until the precipitate just dissolves.
  4. Dilute to 100 mL with distilled water.

Tissue Sample

5 µ paraffin sections of neutral buffered formalin fixed 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 2% silver nitrate for 24 hours.
  3. Treat with ammoniacal silver solution for 20-40 minutes, until they are brown.
  4. Rinse with distilled water.
  5. Place in 12% formalin for 5-10 minutes.
  6. Wash well with tap water.
  7. Rinse with distilled water.
  8. Tone with 0.5% gold chloride solution for 2 hours.
  9. Rinse well with distilled water.
  10. Fix in 5% sodium thiosulphate for 10-15 minutes.
  11. Wash well with tap water.
  12. Counterstain with neutral red for 1 minute.
  13. Rinse with tap water.
  14. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Nuclei  –  red
  • Background  –  grey

Notes

  • Although the instructions do not specify it, any mercury pigment from the Zenker’s fluid fixation may be removed with the iodine-thiosulphate sequence.
  • 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.
  • 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.
March 30, 2023
Love0

Lillie’s Impregnation for Reticulin

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

Lillie's Impregnation

for Reticulin

20
steps
8
materials
print

Materials

  • Silver nitrate, 10% aqu.
  • Strong ammonium hydroxide (s.g. 0.88).
  • Potassium permanganate, 0.5% aqu.
  • Oxalic acid, 5% aqu.
  • Uranium nitrate, 1% aqu.
  • Formalin, 10% aqu.
  • Yellow gold chloride, 0.2% aqu.
  • Sodium thiosulphate, 5% aqu.
  • Neutral red, 1% aqu.

Preparation of Lillie’s Ammoniacal Silver

  1. Place 20 mL strong ammonium hydroxide in a flask.
  2. Add 10% silver nitrate drop by drop until the precipitate is almost dissolved (about 25-30 mL). The solution should have a faint opalescence.
  3. Dilute to twice its volume with distilled water.

Tissue Sample

5 µ paraffin sections of neutral buffered formalin fixed 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. Oxidise with 0.5% potassium permanganate for 2 minutes.
  3. Rinse well with tap water.
  4. Bleach in Oxalic acid for a 2 minutes.
  5. Rinse well with tap water.
  6. Sensitise with 1% uranium nitrate solution for 5-10 seconds.
  7. Rinse with distilled water.
  8. Treat with ammoniacal silver solution for 3 minute.
  9. Rinse briefly with 90% ethanol.
  10. Rinse with distilled water.
  11. Place in 10% formalin for 2 minutes.
  12. Rinse well with tap water.
  13. Rinse with distilled water.
  14. Tone with 0.2% gold chloride solution.
  15. Rinse with distilled water.
  16. Fix in 5% sodium thiosulphate for 2 minute.
  17. Wash well with running tap water.
  18. Counterstain with neutral red for 1 minute.
  19. Rinse with tap water.
  20. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Nuclei  –  red
  • Background  –  grey

Notes

  • 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.
  • It is sometimes difficult to obtain uranium nitrate, particularly if it requires international transportation.
  • 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.
March 30, 2023
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Hitchcock Ehrich for Plasma Cells

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

Hitchcock Ehrich

for Plasma Cells

5

steps
3

materials
print

This method uses malachite green and acridine red to stain plasma cells in a similar manner to the methyl green pyronin methods.


Materials

Solution A

MaterialAmount
Malachite green1g
Distilled water100mL

Solution B

MaterialAmount
Acridine red3g
Distilled water100mL

Combine 1 part of solution A with 3 parts of solution B immediately before use.

Tissue Sample

Zenker fixation is recommended. Other fixatives may not be satisfactory.


Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into the staining solution for 30 seconds.
  3. Rinse with water.
  4. Dehydrate rapidly with absolute ethanol.
  5. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  green
  • Plasma cell cytoplasm  –  crimson
  • 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. Biological Staining Methods, 6th ed. (1957)
    Gurr George T.,
    George T. Gurr, London, UK
March 30, 2023
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