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Aldehyde Toluidine Blue for Mast Cells

By Intracytoplasmic Granules, Mast Cells, Protocols, Stain Target

Aldehyde Toluidine Blue

for Mast Cells

7
steps
6
materials

Materials

Compounding Procedure

  1. Dissolve the dye in the ethanol.
  2. Add paraldehyde and acid.
  3. Ripen one week at room temperature.
  4. Store at room temperature.
  5. Filter before use. Stable for a year or longer.

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. Rinse with 70% ethanol.
  3. Place in solution A for 1 hour.
  4. Wash off excess stain with 70% ethanol.
  5. Rinse well with tap water.
  6. Counterstain with nuclear fast red-tartrazine.
  7. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Mast cell granules  –  deep blue
  • Nuclei  –  red
  • Background  –  yellow

Notes

  • The staining solution is a modification of Gomori’s aldehyde fuchsin using toluidine blue instead of basic fuchsin.
  • Staining time may need to be increased as the solution ages (up to 2 hours). If staining takes longer than 2 hours, prepare a new solution.
  • Elastic fibres are unstained, likely because basic fuchsin can form dipole-dipole interactions and toluidine blue generally does not.
  • Mucins are stained very pale blue.
  • Different samples of this dye may vary in effectiveness. If a sample gives pale staining, try one from another vendor. Toluidine blue from Fisher Scientific was used to develop the method.

Safety Note

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

References

  1. Llewellyn, B. D., unpublished.

Naphthoic Acid Hydrazide for Aldehydes

By Aldehydes, Protocols, Stain Target

Naphthoic Acid Hydrazide

for Aldehydes

11
steps
11
materials

Materials

Veronal acetate

MaterialAmount
Sodium acetate1.943g
Sodium barbiturate2.943g
Distilled waterto 100mL

Veronal buffer pH 7.4

MaterialAmount
Veronal acetate solution5mL
M/10 hydrochloric acid5mL
Distilled water60mL

NAH

MaterialAmount
2-hydroxy-3-naphthoic acid hydrazide0.1g
Ethanol, 100%95mL
Acetic acid, glacial5mL

Fast blue B

MaterialAmount
Fast blue B salt0.1g
Veronal acetate buffer (pH 7.4)100mL

Tissue Sample

5 µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Many other fixatives are satisfactory. Fixatives containing strong acids should be avoided if the intent is to demonstrate aldehydes generated from acid hydrolysis of DNA, as acids in some fixatives may hydrolyse the tissue during fixation (picric acid in Bouin’s formal-picric-acetic mixture, for example).

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Oxidise or hydrolyse to generate aldehydes.
  3. Rinse briefly with distilled water.
  4. Rinse briefly with 50% ethanol.
  5. Place into NAH solution at room temperature for 3-6 hours.
  6. Rinse with three changes of 50% ethanol, about 10 minutes each.
  7. Wash well with water.
  8. Place into pre-cooled fast blue B solution for 1-3 minutes at 0°C.
  9. Wash well with water.
  10. Optionally, counterstain appropriately.
  11. Dehydrate with ethanol, clear with xylene, and mount with a resinous medium.

Expected Results

  • Aldehyde sites  –  blue to purple
  • Background  –  as counterstained

Notes

  • Sodium barbiturate is also known as veronal.
  • Procedures for producing aldehydes include those for acid hydrolysis of DNA, periodic acid and chromic acid oxidation of carbohydrates. In those procedures, begin at step 3, above, where those other methods specify placing into Schiff’s reagent.

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).
    Cellular pathology technique., Ed. 4., p. 187
    Butterworths, London, England.

Cook & Lamb Alkali Blue for Elastic Fibres

By Elastic Fibers, Protocols, Stain Target

Cook & Lamb Alkali Blue

for Elastic Fibres

10
steps
3
materials

Materials

  • 0.4% alkali blue in 70% ethanol
  • 4% aqueous ferric ammonium sulphate
  • 0.05% potassium hydroxide in absolute ethanol

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. Place in alkali blue solution for 5 minutes.
  3. Wash with water.
  4. Place in iron alum solution for 3 minutes.
  5. Wash with water.
  6. Rinse with ethanol.
  7. Rinse briefly with alcoholic potassium hydroxide to clear the background.
  8. Counterstain if wished.
  9. Dehydrate with ethanols.
  10. Clear with xylene and mount with a resinous medium.

Expected Results

  • Elastic fibres – blue (blue-green after Van Gieson)
  • Erythrocytes – pale blue-green

Notes

  • Neutral red, safranin, hemalum and eosin are suggested as counterstains.
  • The alkali blue solution lasts for several weeks.
  • This is an example of afterchrome mordanting.

Safety Note

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

References

  1. Histological demonstration techniques, (1974))
    Cook, H C.
    Butterworths, London, England

Allen’s Stain for Mast Cells

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

Allen's Stain

for Mast Cells

11
steps
3
materials

Materials

Tissue Sample

Paraffin sections of neutral buffered formalin fixed tissue are suitable. Mercuric chloride fixatives are reputed to emphasise metachromasia. Other fixatives may be satisfactory.


Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Rinse with water.
  3. Stain nuclei lightly with alum hematoxylin.
  4. Rinse with tap water and blue hematoxylin.
  5. Rinse well with water.
  6. Place in neutral red for 10 minutes.
  7. Rinse with distilled water.
  8. Differentiate with 70% ethanol up to 10 minutes.
  9. Dehydrate with 96% ethanol up to 5 minutes.
  10. Dehydrate with N-butanol up to 10 minutes.
  11. Clear with xylene and mount using a resinous medium.

Expected Results

  • Nuclei – blue
  • Mast cell granules = red

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, (1976).
    Lynch’s Medical Laboratory Technology., Ed. 3. Vol. II. pp. 980
    W. B. Saunders Company, Toronto, Canada.
  2. Putt, F A, (1972).
    Manual of Histopathological Staining Methods., pp. 233
    John Wiley & Sons, London, UK.

Lillie’s Allochrome

By Periodic Acid-Schiff Reaction, Protocols, Schiff's Reagent Reactions, Stain Type
Protocol

Lillie's Allochrome

8
steps
5
materials

Materials

  • 1% Aqueous periodic acid
  • Schiff’s reagent
  • Weigert’s iron hematoxylin
  • Picro-methyl blue
    MaterialAmount
    Saturated aqueous picric acid100mL
    1% aqueous methyl blue0.4mL

Tissue Sample

5 µ paraffin sections. Most fixatives are satisfactory, including 10% NBF, but those generally considered preferable for trichrome stains will be advantageous.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Do a PAS stain, but do not counterstain.
  3. Stain nuclei with Weigert’s iron hematoxylin or equivalent.
  4. Wash well in tap water, rinse with distilled water.
  5. Place into picro-methyl blue for 6 minutes.
  6. Rinse well with 95% ethanol.
  7. Dehydrate with absolute ethanol.
  8. Clear with xylene and mount with a resinous medium.

Expected Results

  • Nuclei – Brown-black
  • Collagen & reticulin – blue
  • Cytoplasm & muscle – yellow
  • PAS positive structures – red

Safety Note

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

References

  1. Histological demonstration techniques, (1974)
    Cook, H C.
    Butterworths, London, England

Bennhold’s Congo Red for Amyloid

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

Bennhold's Congo Red

for Amyloid

8
steps
5
materials

Materials

  • Mayer’s hemalum
  • Congo red
    MaterialAmount
    Congo red1g
    Distilled water100mL
  • Lithium carbonate
    MaterialAmount
    Lithium carbonateasrequired
    Distilled water100mL

Dissolve to saturation.

Tissue Sample

Paraffin sections of formalin fixed tissues are satisfactory.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into the congo red solution for
    1. 30 minutes at room temperature, or
    2. 45 minutes at 56°C, or
    3. 15 seconds at boiling.
  3. Drain and, without rinsing, place into lithium carbonate solution for 15 seconds.
  4. Drain and, without rinsing, differentiate in 80% ethanol. This usually takes just a few seconds.
  5. Wash well with tap water.
  6. Stain nuclei with hematoxylin and blue.
  7. Dehydrate with ethanol.
  8. Clear with xylene and mount with a resinous medium.

Expected Results

  • Amyloid  –  red
  • Nuclei  –  blue

Notes

  • Amyloid displays apple green birefringence when viewed with crossed polarisers, one above and one below the section.
  • The congo red is most commonly applied for 30 minutes at room temperature.
  • Differentiation in 80% ethanol is difficult to control, and amyloid is often poorly demonstrated. Due to this the method is not usually recommended.

Safety Note

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

References

  1. Bennhold, H., (1922),
    Eine specifische amyloidfärbung mit kongorot,
    Münchener Medizinische Wochenschrift, v 44, page 1537

Bencosme’s HPS General Oversight stain

By Protocols, Stain Type, Trichrome Staining, Yellowsolve Staining

Bencosme's HPS

General Oversight stain

11
steps
5
materials

Also known as the Hematoxylin Phloxine Saffron stain.

Materials

  • Regressive hemalum
  • Solution A
    MaterialAmount
    Phloxine B2g
    Distilled water100mL

    Dissolve the phloxine B and filter. Preserve with 2-5 drops of strong formalin.

  • Solution B
    MaterialAmount
    Saffron6g
    Absolute ethanol200mL

    Mix the saffron stigmata with the ethanol and seal tightly. Place in an oven at 56°C for 1-2 weeks.

Tissue Sample

Brasil’s fixative was specified. 5 µ paraffin sections of neutral buffered formalin fixed tissue are suitable, but results may be improved by refixing in Bouin’s fixative or saturated aqueous picric acid at 56°C for one hour prior to staining, then washing the sections in tap water to remove all yellow discoloration.

Protocol

  1. Bring sections to water with xylene and ethanol.
  2. Treat with Bouin’s fluid if desired.
  3. Stain nuclei with hemalum, differentiate and blue.
  4. Wash well with water.
  5. Place in solution A for 2-20 minutes.
  6. Wash with tap water until red stain stops being removed, 1-5 minutes.
  7. Differentiate phloxine with 80% ethanol up to 1½ minutes.
  8. Thoroughly dehydrate with absolute ethanol.
  9. Place into solution B for 2-20 minutes until a suitable red-yellow balance is achieved.
  10. Rinse well with absolute ethanol.
  11. Clear with xylene and mount with a synthetic resinous medium.

Expected Results

  • Nuclei  –  blue
  • Cytoplasm  –  red shades
  • Muscle  –  pink
  • Elastic fibres  –  bright red
  • Collagen  –  orange-yellow

Notes

  • Any hemalum may be used, but Bencosme specified a particular formula.
  • Saffron is expensive. It may be available in East Indian grocery stores or health food stores as its most common use today is as a spice and food coloring. Usually, whole stigmata are more effective than ground saffron.
  • If the saffron solution is too strong it may be diluted with absolute ethanol. Store and use at room temperature. It has a limited life of about a month and is best when freshly made. It is important that there be no water in the saffron solution and that sections be thoroughly dehydrated before it is applied. If contaminated with moisture the solution must be discarded.

Safety Note

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

References

  1. Medical Laboratory Technology, 2nd ed. (1969)
    Lynch M.J., Raphael S.S., Mellor L.D., Spare P.D. and Inwood M.J.H.,
    W. B. Saunders Co., Toronto, On., Canada

Congo Red Fluorescence for Amyloid

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

Congo Red Fluorescence

for Amyloid

8
steps
4
materials

Expected Results

  • Amyloid  –  Orange to red fluorescence

Materials

Congo red

MaterialAmount
Congo red0.1g
Ethanol, 50%100mL

Differentiator

MaterialAmount
Potassium hydroxide0.2g
Ethanol, 80%100mL

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 congo red for 1 minute.
  3. Wash with tap water.
  4. Place into differentiator until the section appears to be unstained.
  5. Wash with water.
  6. Dehydrate with absolute ethanol.
  7. Clear with xylene.
  8. Mount with a fluorescence free resinous medium.

Notes

  • This method is very similar to Highmans procedure.

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.

Eastwood and Cole Congo Red for Amyloid

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

Eastwood and Cole Congo Red

for Amyloid

6
steps
7
materials

Materials

  • Mayer’s hemalum
  • Buffer pH 10
    MaterialAmount
    Glycine, 0.1M30mL
    Sodium chloride, 0.1M30mL
    Sodium hydroxide, 0.1M40mL
  • Congo red
    MaterialAmount
    Congo red0.5g
    Buffer pH 1050mL
    Ethanol, absolute50mL

    Combine the ethanol and buffer. Dissolve the congo red.

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 hemalum and blue.
  3. Place in congo red solution for 10 – 20 minutes.
  4. Rinse off the staining solution with 70% ethanol until clear.
  5. Dehydrate with ethanol.
  6. Clear with xylene and mount with a resinous medium.

Expected Results

  • Amyloid  –  Orange to red
  • Eosinophils and elastic  –  Orange to red
  • Nuclei  –  Blue

Notes

  • Amyloid displays apple green birefringence when viewed with crossed polarisers, 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. Bancroft, J.D. and Stevens A. (1982)
    Theory and practice of histological techniques Ed. 2
    Churchill Livingstone, Edinburgh & London, UK.

Highman’s Congo Red for Amyloid

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

Highman's Congo Red

for Amyloid

7
steps
6
materials

Materials

  • Mayer’s hemalum
  • Congo red
    MaterialAmount
    Congo red0.5g
    Distilled water50mL
    Ethanol, 100%50mL
  • Alkaline ethanol
    MaterialAmount
    Ethanol, 80%100mL
    Potassium hydroxide0.2g

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. Place into congo red solution for 5 minutes or longer.
  3. Differentiate in alkaline ethanol (about 5-30 seconds).
  4. Wash well with tap water.
  5. Stain nuclei with hematoxylin and blue.
  6. Dehydrate with ethanol.
  7. Clear with xylene and mount with a resinous medium.

Expected Results

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

Notes

  • Amyloid displays apple green birefringence when viewed with crossed polarisers, one above and one below the section.
  • Sirius red F3B may also be used in this method.

Safety Note

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

References

  1. Highman, B., (1946),
    Improved methods for demonstrating amyloid in paraffin sections,
    Archives of Pathology, v 41, page 559
  2. Bancroft, J. D. and Stevens, A.
    Theory and practice of histological techniques,
    Churchill Livingstone, London, England