Category

Stain Target

Lendrum’s Phloxine Tartrazine for Viral Inclusion Bodies

By Intracytoplasmic Granules, Paneth Cells, Protocols, Stain Target, Stain Type, Trichrome Staining, Yellowsolve Staining
Protocol

Lendrum's Phloxine Tartrazine

for Viral Inclusion Bodies

11
steps
6
materials
print

This method is also used for the demonstration of Paneth cell granules, and may be used as a substitute for the HPS if the differentiation in the tartrazine solution is shortened to retain pink cytoplasm and muscle.

Materials

  • Mayer’s hemalum
  • Solution A
    MaterialAmount
    Phloxine B0.5g
    Calcium chloride0.5g
    Distilled water100mL
  • Solution B
    MaterialAmount
    Tartrazineto saturation
    2-Ethoxy ethanol (cellosolve)100mL

Tissue Sample

5µ paraffin sections of formal sublimate fixed tissue is preferred. Formalin fixed tissue is suitable. Other fixatives are likely to be satisfactory.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Stain nuclei to medium density with hemalum.
  3. Wash in tap water for 5 minutes. Blueing takes place in the phloxine solution.
  4. Place in solution A for 20 minutes.
  5. Rinse in tap water, blot almost dry. Some technologists rinse with cellosolve instead of blotting. The object is to remove all traces of water, as it interferes with the ability of tartrazine to extract phloxine and counterstain.
  6. Rinse with solution B to remove remaining water. Discard solution.
  7. Place in solution B until inclusions are red and all other tissue is yellow. The time varies considerably. Control microscopically.
  8. Rinse thoroughly but briefly with absolute ethanol.
    Do not rinse with water at this stage as it rapidly removes the tartrazine. Some technologists use cellosolve instead of ethanol.
  9. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei  –  blue
  • Acidophil virus inclusion bodies  –  red
  • Paneth cell granules  –  red
  • Background  –  yellow

Notes

  • The 2-ethoxy ethanol must not be replaced by any other solvent.
  • The 2-ethoxy ethanol must be, and must remain, completely anhydrous.

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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Jalowy’s Impregnation for Reticulin

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

Jalowy's Impregnation

for Reticulin

8
steps
4
materials
print

Materials

  • Silver nitrate, 10% aqu.
  • Strong ammonium hydroxide (s.g. 0.88).
  • Ammonium hydroxide, 1% aqu.
  • Formalin, 10% aqu.

Preparation of Jalowy’s Ammoniacal Silver

  1. Place 20 mL of 10% silver nitrate in a flask and add 1 mL of 40% sodium hydroxide.
  2. Mix well, then filter. Wash the precipitate well with distilled water several times and decant.
  3. Add 20 mL distilled water to the precipitate and suspend.
  4. Add strong ammonium hydroxide by drops until the precipitate is just dissolved.
  5. 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. Rinse with distilled water.
  3. Place in prewarmed ammoniacal silver solution for 5-30 minutes at 30°C.
  4. Rinse with distilled water.
  5. Rinse with 1% ammonium hydroxide.
  6. Place in 10% formalin for 2-10 minutes.
  7. Rinse well with tap water.
  8. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Reticulin fibres  –  black
  • Background  –  brown

Notes

  • This method was recommended for collagen and reticulin fibres on formalin fixed skin.
  • Ensure that the ammonium hydroxide and sodium hydroxide are fresh and full strength. Keep the ammonium hydroxide well stoppered when not in use. Pour sufficient for use into a beaker, then immediately restopper the stock container. Do not return unused ammonium hydroxide to the stock bottle.
  • The instructions for this method do not specify to tone, fix, or counterstain. If fading or a lack of contrast make it desirable, toning with 0.1% yellow gold chloride until satisfactory, and/or fixing with 5% sodium thiosulphate, and/or counterstaining with 1% aqueous neutral red may be used.

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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Kurnick Stain for Plasma Cells

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

Kurnick Stain

for Plasma Cells

5
steps
3
materials
print

Materials

Stock pyronin

MaterialAmount
Pyronin Y2g
Distilled water100mL

Stock methyl green

MaterialAmount
Methyl green2g
Distilled water100mL

Working solution

MaterialAmount
Stock pyronin12.5mL
Stock methyl green7.5mL
Distilled water30mL

Storing stock solutions

The pyronin stock solution should be extracted with chloroform ten times to ensure purity. The methyl green stock solution should be extracted a minimum of six times, until the chloroform is no longer colored with any crystal violet. Store both solutions over a layer of chloroform.

Tissue Sample

Most fixatives should be satisfactory if fixation time is not extended, but avoid potassium dichromate or picric acid. Overnight with a 10% formalin variant should be satisfactory.

Protocol

  1. Bring sections to distilled water via xylene and ethanol.
  2. Place into the staining solution for 6 minutes in a coplin jar.
  3. Blot gently.
  4. Dehydrate with n-butanol, two changes of 5 minutes each.
  5. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei – green to blue-green
  • Plasma cell cytoplasm, Nissl substance, nucleoli – red
  • Other cell cytoplasm – pink to unstained

Notes

  • As with other methyl green-pyronin methods used for demonstrating nucleic acids, in critical applications a control section should be treated with a 0.1% solution of ribonuclease for one hour at 37°C to remove RNA, and another treated with distilled water alone. Both control sections should be compared with an untreated but stained section.

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. pp.187-188.
    E. & S. Livingstone, Edinburgh and London, UK.
March 30, 2023
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Romanowsky – Giemsa General Oversight Stain

By Eosinophils, Intracytoplasmic Granules, Protocols, Stain Target
Protocol

Romanowsky – Giemsa

General Oversight Stain

9
steps
3
materials
print

Materials

  • Stock Giemsa or other Romanowsky stain e.g. Lieshman, Wright etc.
  • 0.1% acetic acid in distilled water.

Working Solution

MaterialAmount
Stock Romanowsky stain1mL
Distilled water or pH 6.8 buffer9mL

Make the diluted solution just before using. Discard after a few hours.

Tissue Sample

Most fixatives permit staining but the results may vary. 3 µ paraffin sections of neutral buffered formalin fixed tissue are usually suitable.

Protocol

  1. Bring sections to water with xylene and ethanol.
  2. Optionally, treat with pH 6.8 phosphate buffer for 30 minutes.
  3. Place into the staining solution for 1 hour.
  4. Rinse well with water.
  5. Differentiate with acetic acid until nuclear morphology is clear. Control microscopically.
  6. Rinse well with distilled water.
  7. Blot dry with filter paper, then flood with xylene.
  8. Repeat step 6 until section is transparent (usually 4-5 times).
  9. Mount with a synthetic resinous medium.

Expected Results

  • Nuclei  –  blue
  • Cytoplasm  –  pink

Notes

  • Giemsa is usually diluted 1 in 10. Lieshman, Wright and others are often diluted 1 in 3.
  • Pretreatment with a pH 6.8 phosphate buffer is sometimes recommended immediately before placing in diluted stain. In that case, buffer with the same pH should be used to dilute the stock Romanowsky stain.
  • Differentiation may also be carried out in the same buffer as used to dilute the stock stain, but may take some time.
  • Drury and Wallington use pH 5.0 buffer mixed equal parts with methanol, and specify green Euparal as the mounting medium.
  • Sections may be rapidly dehydrated with ethanol instead of blotting, but this removes some blue staining and will likely destroy any metachromasia.
  • Sections stained with Romanowsky stains do not usually show the same range of colors that are shown by the same stain on smears, and the choice of stock Romanowsky stain does not necessarily influence the final appearance.

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., Alison, R.T. and Barr, W.T. (1985)
    Cellular Pathology Technique, 4th ed.
    Butterworths, London, UK.
  2. 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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Gluck’s Impregnation for Reticulin on Zenker Fixed Tissue

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

Gluck's Impregnation

for Reticulin on Zenker Fixed Tissue

20
steps
12
materials
print

Materials

  • Oxalic acid, 5% aqu.
  • Silver nitrate, 10% aqu.
  • Strong ammonium hydroxide (s.g. 0.88).
  • Sodium hydroxide, 40% aqu.
  • Formalin, 5% aqu.
  • Yellow gold chloride, 0.2% aqu.
  • Neutral red, 1% aqu.
  • Gram’s iodine.
    MaterialAmount
    Iodine1g
    Potassium iodide2g
    Distilled water300mL

    Mix the iodine and potassium iodide in a 500 mL flask. Add 5 mL of the water. When the iodine has dissolved, make up to 300 mL with distilled water.

  • Cajal’s Solution
    MaterialAmount
    Strong formalin15mL
    Ammonium bromide2g
    Water85mL

Preparation of Gluck’s Ammoniacal Silver

  1. Place 10 mL of 10% silver nitrate in a flask.
  2. Add 0.5 mL of 40% sodium hydroxide.
  3. Allow to settle, then decant the supernatent.
  4. Wash the precipitate, allow to settle, then decant a few times.
  5. While swirling, slowly add drops of strong ammonium hydroxide until the precipitate just redissolves.
  6. Dilute to 100 mL with distilled water, then add 2 mL pyridine.

Tissue Sample

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

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Rinse well with tap water.
  3. Place in Gram’s iodine for 4-8 hours.
  4. Rinsewith 70% ethanol.
  5. Rinse well with tap water.
  6. Bleach in 5% sodium thiosulphate for 5 minutes.
  7. Wash well with tap water.
  8. Place in Cajal’s solution at 37°C for 24 hours.
  9. Wash with tap water.
  10. Place in Gluck’s ammoniacal silver for 5 minutes.
  11. Rinse with distilled water.
  12. Place in 5% formalin for 5 minutes.
  13. Wash with tap water.
  14. Tone with 0.2% gold chloride solution until grey.
  15. Rinse with distilled water.
  16. Place in 5% sodium thiosulphate for 5 minutes.
  17. Wash well with 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 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 but before adding the pyridine, 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.
  • The formalin used to make Cajal’s solution 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. 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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Gridley’s Impregnation for Reticulin

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

Gridley's Impregnation

for Reticulin

17
steps
9
materials
print

Materials

  • Silver nitrate, 2% aqu.
  • Silver nitrate, 20% aqu.
  • Strong ammonium hydroxide (s.g. 0.88).
  • Sodium hydroxide, 40% aqu.
  • Periodic acid, 0.5% aqu.
  • Formalin, 3% aqu.
  • Yellow gold chloride, 0.5% aqu.
  • Sodium thiosulphate, 5% aqu.
  • Neutral red, 1% aqu.

Preparation of Da Fano’s Ammoniacal Silver

  1. Place 10 mL of 20% silver nitrate in a flask.
  2. Add 0.2 mL of 40% sodium hydroxide.
  3. While swirling, slowly add drops of strong ammonium hydroxide until the precipitate just redissolves.
  4. Make up to 80 mL with distilled water.

Tissue Sample

6 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% periodic acid for 15 minutes.
  3. Rinse well with distilled water.
  4. Sensitise with 2% silver nitrate for 30 minutes.
  5. Rinse well with distilled water.
  6. Treat with da Fano’s ammoniacal silver for 15 minutes.
  7. Rinse with distilled water.
  8. Reduce in 3% formalin for 3 minutes.
  9. Rinse well with tap water.
  10. Rinse with distilled water.
  11. Tone with 0.5% gold chloride solution for 5 minutes.
  12. Rinse with distilled water.
  13. Fix in 5% sodium thiosulphate for 5 minutes.
  14. Wash well with running tap water.
  15. Counterstain with neutral red for 1 minute.
  16. Rinse with tap water.
  17. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

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

Notes

  • Ensure that both the ammonium hydroxide and potassium 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.
  • 3% formalin is made by diluting strong formalin with tap water (3 mL strong formalin, 97 mL tap water).
  • Toning is a variable step. Untoned sections give dark brown reticulin fibres on a paler brown background. Although the method specifies 5 minutes toning in 0.5% gold chloride, this is not mandatory. 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.
  • The use of periodic acid followed by an easily reducible silver solution is obviously similar to the Jones’ method for basement membranes and carbohydrates. Note, however, that Gridley’s method applies ammoniacal silver solution at room temperature after sensitising in aqueous silver nitrate at room temperature, then applies formalin as a reducing agent, i.e. it is an argyrophil reaction, while the Jones’ method applies a methenamine silver solution at 56°C for a longer time, without sensitising, and does not use an external reducer, i.e. it is an induced argentaffin reaction. Clearly, in Gridley’s method the periodic acid is being used for the same purpose as the potassium permanganate of the Mallory bleach in other methods.

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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Gordon & Sweets Impregnation for Reticulin

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

Gordon & Sweets Impregnation

for Reticulin

19
steps
11
materials
print

Materials

  • Silver nitrate, 10% aqu.
  • Strong ammonium hydroxide (s.g. 0.880).
  • Sodium hydroxide, 3% aqu.
  • Oxalic acid, 1% aqu.
  • Iron alum, 2.5% aqu.
  • Formalin, 10% aqu.
  • Yellow gold chloride, 0.2% aqu.
  • Sodium thiosulphate, 3% aqu.
  • Neutral red, 1% aqu.
  • Mallory bleach
    MaterialAmount
    Potassium permanganate, 0.5% aqu.47.5mL
    Sulphuric acid, 3% aqueous2.5mL

Preparation of Ammoniacal Silver

  1. Place 5 mL of the 10% silver nitrate in a 100 mL flask.
  2. Using a pasteur pipette, add a drop of strong ammonium hydroxide then swirl the solution for a few seconds.
  3. A precipitate will form at first.
  4. Continue adding ammonium hydroxide drop by drop and swirling until the precipitate is just redissolved.
  5. Add 5 mL of 3% sodium hydroxide and mix well.
  6. A precipitate will again form.
  7. Add drops of ammonium hydroxide until the precipitate is just redissolved, leaving a faint opalescence.
  8. Dilute to 50 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. Oxidise with the Mallory bleach for 5 minutes.
  3. Bleach in Oxalic acid for a few minutes.
  4. Rinse with distilled water.
  5. Rinse well with tap water.
  6. Sensitise with iron alum solution for 15 minutes.
  7. Rinse well with distilled water.
  8. Treat with ammoniacal silver for 30 seconds.
  9. Rinse well with distilled water.
  10. Reduce in formalin 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 sodium thiosulphate for 10 minutes.
  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

  • Gordon & Sweets also suggested a variant that used one of Foot’s ammoniacal silver solutions. Apart from that, the method is the same as given above.

    Foot’s ammoniacal silver

    • To 10 mL of 1% silver nitrate, add 0.1 mL of 40% potassium hydroxide.
    • Add strong ammonium hydroxide by drops until the precipitate just redissolves.
    • Make up the volume to 100mL 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.
  • Most references to the Gordon & Sweets’ reticulin stain specify that the ammoniacal silver solution should be made with 10.2% aqueous silver nitrate and 3.1% aqueous sodium hydroxide. No explanation is given. The 10% and 3% solutions respectively, as given by Gray, work satisfactorily.
  • Iron alum is ferric ammonium sulphate. For routine formalin fixed tissue 15 minutes in the iron alum is usually sufficient. If necessary the time may be extended up to 2 hours.
  • 10% formalin is made by diluting strong formalin 1:10 with tap water (10 mL strong formalin, 90 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. Drury, R A, and Wallington, E A, (1967).
    Carleton’s histological technique., Ed. 5.
    Oxford University Press, London, England.
  2. Culling, C F A, Allison, R T, Barr, W T, (1985).
    Cellular pathology technique., Ed. 4.
    Butterworths, London, England.
  3. 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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Slidders’ Fuchsin-Miller for Fibrin

By Fibrin, Protocols, Stain Target, Stain Type, Trichrome Staining, Yellowsolve Staining
Protocol

Slidders’ Fuchsin-Miller

for Fibrin

16
steps
7
materials
print

Materials

  • An acid resistant nuclear stain, such as Weigert’s iron hematoxylin, or the celestine blue-hemalum sequence.
  • Fuchsin
    MaterialAmount
    Acid fuchsin1g
    Acetic acid, glacial2.5mL
    Distilled water100mL
  • Miller
    MaterialAmount
    Milling yellow 3G2.5g
    2-ethoxy-ethanol100mL
  • Phosphotungstic acid
    MaterialAmount
    Phosphotungstic acid1g
    Distilled water100mL

Tissue Sample

3 mm slices of tissue should be fixed in formol sublimate or B5 overnight. Paraffin process overnight. Overnight formalin fixation and paraffin processing can produce acceptable results if sections are refixed for an hour in Bouin’s picro-acetic-formalin mixture at 56°C. Avoid rapid fixation with formalin and short processing, as this produces tissues that stain poorly even with the secondary fixation specified. Optimal results are obtained with extended mercuric chloride fixation, thorough processing, degreasing and secondary fixation of sections. Sections should be 3-5 µ thick.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. If mercury fixed, remove with the iodine-thiosulphate sequence.
  3. If not mercury fixed, place into Bouin’s fluid at 56°C for 1 hour.
  4. Rinse well with distilled water.
  5. Stain nuclei with an acid resistant nuclear stain.
  6. Rinse with 95% ethanol.
  7. Place in fuchsin for 10 minutes.
  8. Rinse with distilled water.
  9. Differentiate with phosphotungstic acid for 5 minutes.
  10. Rinse well with distilled water.
  11. Blot.
  12. Rinse well with cellosolve.
  13. Place into milling yellow until fibrin is red and tissues are yellow.
    This step takes from ½ – 4 hours.
  14. Rinse briefly with 1% aqueous acetic acid.
  15. Rinse with cellosolve.
  16. Clear with xylene and mount with a resinous medium.

Expected Results

  • Fibrin  –  red
  • Other tissue  –  yellow
  • Nuclei  –  black

Notes

  • It is very important that the milling yellow solution be completely anhydrous. Even small amounts of water or other solvents can cause problems with displacement and incomplete removal of red dye. For that reason, step 12 should be thorough and the milling yellow should be used in a Coplin jar with a lid sealed with tape.
  • A well stained section would show red fibrin only, muscle and erythrocytes being yellow. With poorly fixed material, both erythrocytes and muscle fibres may resist displacement of the red dye and they may have some residual red colouring. Sometimes this may be overcome by prestaining either with saturated picric acid in absolute ethanol or the MSB martius yellow solution for two minutes immediately before step 7, but a better resolution is correct fixation and processing.
  • Some intracellular materials may stain red, such as Paneth cell granules.

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.
  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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Martius, Scarlet and Blue (MSB) for Fibrin

By Fibrin, Protocols, Stain Target, Stain Type, Trichrome Staining, Trichrome, Multi-Step
Protocol

Martius, Scarlet and Blue (MSB)

for Fibrin

14
steps
8
materials
print

The MSB (Martius, Scarlet and Blue) method for fibrin is a reliable technique. It is more automatic than other methods, i.e. it is less dependent on skill and experience and is consequently suitable for a routine laboratory. Overnight mercuric chloride fixation (formol sublimate or B5) is preferred, followed by overnight paraffin processing, although formalin fixed, paraffin embedded material can produce acceptable results if sections are refixed for an hour in Bouin’s picro-acetic-formalin mixture at 56°C. Optimal results are obtained with extended mercuric chloride fixation, thorough processing, degreasing and secondary fixation of sections, as for the Picro-Mallory.

Materials

  • An acid resistant nuclear stain, such as Weigert’s iron hematoxylin, or the celestine blue-hemalum sequence.
  • Martius
    MaterialAmount
    Martius yellow0.5g
    Phosphotungstic acid2g
    Ethanol, 95%100mL
  • Scarlet
    MaterialAmount
    Crystal scarlet1g
    Acetic acid, glacial2.5mL
    Distilled water97.5mL
  • Blue
    MaterialAmount
    Methyl blue0.5g
    Acetic acid, glacial1mL
    Distilled water98mL
  • Phosphotungstic acid
    MaterialAmount
    Phosphotungstic acid1g
    Distilled water100mL

Tissue Sample

3 mm slices of tissue should be fixed in formol sublimate (or B5) overnight. Paraffin process overnight. Overnight formalin fixation is usually satisfactory, but avoid rapid fixation with formalin and short processing, as this produces tissues that stain poorly even with the secondary fixation specified. Sections should be 3-5 µ thick.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. If mercury fixed, remove with the iodine-thiosulphate sequence.
  3. If not mercury fixed, place into Bouin’s fluid at 56°C for 1 hour.
  4. Rinse well in distilled water.
  5. Stain nuclei with an acid resistant nuclear stain.
  6. Rinse with 95% ethanol.
  7. Place in martius yellow for 2 minutes.
  8. Rinse with distilled water.
  9. Place in crystal scarlet for 10 minutes.
  10. Differentiate with phosphotungstic acid until only fibrin is red (up to 10 minutes).
  11. Place in methyl blue until collagen is blue (up to 10 minutes).
  12. Rinse briefly with 1% aqueous acetic acid.
  13. Dehydrate rapidly with ethanol.
  14. Clear with xylene and mount with a resinous medium.

Expected Results

  • Fibrin  –  red
  • Fresh fibrin  –  yellow
  • Erythrocytes  –  yellow
  • Connective tissue  –  blue

Notes

  • Crystal scarlet is more commonly known as ponceau 6R.
  • Steps 10 & 11 specify the time as up to. These times should be established by inspection, but will generally remain consistent.
  • Lendrum’s recommendation for formalin fixed material was to dewax sections and treat with trichlorethylene in a sealed contained for 48 hours at 56°C, then to refix in absolute ethanol saturated with both picric acid and mercuric chloride for 24 hours before proceeding to step 2. The alternative treatment with Bouin’s fluid given at step 3 is often satisfactory.
  • Bancroft notes that dyes other than those originally given may be used. Some may not be easily available, and no CI numbers were given.
    ColorDye Options
    YellowLissamine fast yellow
    BlueDurazol blue
    Pontamine sky blue
    Fast green FCF
    Naphthalene black 10B
    RedPonceau de xylidine
    Azofuchsin

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.
  2. Culling C.F.A., (1974)
    Handbook of histopathological and histochemical techniques Ed. 3
    Butterworth, London, UK.
  3. 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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Obadiah for Fibrin

By Fibrin, Protocols, Stain Target, Stain Type, Trichrome Staining, Trichrome, Multi-Step
Protocol

Obadiah

for Fibrin

14
steps
11
materials
print

The name of this stain comes from the letters OBDR45, which refer to the dyes used: Orange, Blue, Direct Red 45 (a synonym for one of the red dyes).

With this trichrome stain full and complete fixation is absolutely essential. Minimalist formalin fixation and quick processing will give poorly stained erythrocytes with red or orange tinges instead of the yellow they should have. Lendrum and coworkers specified about seven days fixation in formal sublimate followed by processing thoroughly, sectioning, degreasing with trichlorethylene for 48 hours, then refixing in picro-mercuric-alcohol. Results are usually poor with formalin fixed material, even if treated with Bouin’s fluid at 56°C for an hour or so.

Materials

  • An acid resistant nuclear stain, such as Weigert’s iron hematoxylin, or the celestine blue-hemalum sequence.
  • Picro-mercuric ethanol
    MaterialAmount
    Ethanol, absolute100mL
    Picric acidtosaturation
    Mercuric chloridetosaturation
  • Orange
    MaterialAmount
    Orange G0.5g
    Phosphotungstic acid1g
    Ethanol, 95%100mL
  • Blue
    MaterialAmount
    Naphthalene blue black CS1g
    Acetic acid, glacial1mL
    Distilled water99mL
  • Red – Option 1
    MaterialAmount
    Chicago red2.5g
    Acetic acid, glacial2.5mL
    Distilled water97.5mL
  • Red – Option 2
    MaterialAmount
    Polar brilliant red BN1g
    Acetic acid, glacial1mL
    Distilled water99mL

Tissue Sample

3 mm slices of tissue should be fixed in formol sublimate for one week. They should be paraffin processed with a schedule that thoroughly and completely dehydrates, then thoroughly cleared with xylene and infiltrated with paraffin wax. This process would normally take 48 hours or longer. Err on the side of completeness, and do not attempt to shorten the process. Avoid rapid fixation and overnight processing, as this produces tissues that stain poorly. Sections should be 3-5 µ thick.

Protocol

  1. Dewax sections with xylene.
  2. Place into a sealed container of trichlorethylene at 56°C for 48 hours.
  3. Rinse well with absolute ethanol.
  4. Refix sections in picro-mercuric-ethanol for a minimum of 3 and preferably 24 hours.
  5. Remove mercury pigment with the iodine-thiosulphate sequence.
  6. Stain nuclei with an acid resistant nuclear stain.
  7. Place in the orange stain for 2 minutes.
  8. Rinse with distilled water.
  9. Place in the Blue stain up to 30 minutes.
  10. Differentiate with the polyacid for 5 minutes.
  11. Place in the Red stain 15-20 minutes (polar brilliant red) or 15-30 minutes (chicago red).
  12. Rinse with distilled water.
  13. Dehydrate with ethanol.
  14. Clear with xylene and mount with a resinous medium.

Expected Results

  • Old fibrin  –  black
  • Younger fibrin  –  may be yellow
  • Connective tissue  –  red

Notes

  • Note that this method reverses the usual order and uses a blue stain before the red stain.
  • Lendrum considered that this method demonstrated fibrin that other methods stained as collagen.

Safety Note

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

References

  1. Lendrum, A. C., et. al. (1962)
    Studies on the character and staining of fibrin.
    Journal of clinical pathology, v. 15, p. 401.
March 30, 2023
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