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Aldehydes

Lillie’s Short PAS Reaction for 1-2 Glycols

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

Lillie's Short PAS Reaction

for 1-2 Glycols

8
steps
5
materials
print

The periodic acid Schiff reaction (PAS) is used to demonstrate the presence of 1-2-glycols, and is consequently an important method in the histochemistry of carbohydrates and the histological demonstration of many structures.

Lillie referred to this method as

The Periodic acid, Schiff Sulfite Leucofuchsin Reaction, short variant.

Materials

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory, although glutaraldehyde should be avoided.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Oxidize in periodic acid for ten minutes.
  3. Wash in running water for five minutes.
  4. Place in Schiff’s reagent for 10 minutes.
  5. Transfer to three successive baths of sodium metabisulfite for 1, 2, and 2 minutes respectively.
  6. Wash in running tap water for 5 minutes.
  7. Counterstain with one of the following:
    1. Place in Mayer’s hemalum for 2 minutes. Wash in running tap water and blue.
    2. Place in Weigert’s iron hematoxylin for 2-4 minutes. Decolorise with Pal’s bleach diluted 1:5 with distilled water. Wash in running tap water for 4 minutes.
    3. Place in Weigert’s iron hematoxylin for 6 minutes. Wash for 4 minutes in running tap water. Place in saturated aqueous picric acid for 1 minute.
  8. Dehydrate with ethanol, clear with xylene, and coverslip using a resinous medium.

Expected Results

  • 1-2-glycols  –  red
  • Nuclei  –  blue or black
  • Cytoplasm  –  yellow or unstained

Notes

  • The nuclear counterstain may obscure some positive staining. Keep the application time short.
  • Weigert’s solution should be allowed to ripen for an hour before use.
  • Weigert’s hematoxylin and Pal’s bleach combination gives highly selective nuclear staining.
  • Picric acid differentiates the nuclear stain as well as coloring the background.
  • Glutaraldehyde fixed tissues will have a non-specific positive background staining. This must be blocked before step 2.

Safety Note

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

References

  1. Lillie, R.D., (1954)
    Histopathologic technique and practical histochemistry Ed.2
    Blakiston, New York, USA.
March 30, 2023
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Lillie’s Long PAS for 1-2-Glycols

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

Lillie's Long PAS

for 1-2-Glycols

14
steps
6
materials
print

The periodic acid Schiff reaction (PAS) is used to demonstrate the presence of 1-2-glycols, and is consequently an important method in the histochemistry of carbohydrates and the histological demonstration of many structures.

Lillie referred to this as

“The Periodic acid Leucofuchsin Method in Relation to Various Modifying Procedures”.

He describes its purpose as being

“To indicate at what points and in what sequence various modifying procedures should be introduced”.

Materials

  • A periodic acid solution.
  • A strong Schiff’s reagent.
  • Sodium metabisulfite, 0.5% aqueous, optional.
  • Mayer’s hemalum, or Weigert’s iron hematoxylin.
  • Picric acid, saturated aqueous, optional.
  • Orange G, 2% aqueous, optional.
  • Methyl blue, 0.1% in saturated aqueous picric acid, optional.

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory, although glutaraldehyde should be avoided.

Protocol

  1. Dewax paraffin sections with xylene.
  2. Remove xylene with two changes of absolute ethanol.
  3. Optionally, do one or more of the following procedures in the order given.
    ProcedureComments
    AcetylationAcetylation blocks hydroxyl and amino groups. Cartilage and glycogen are the most resistant. Other carbohydrates are more easily affected.
    BenzoylationSimilar to acetylation, this also blocks hydroxyl and amino groups. Again, cartilage and glycogen are the most resistant while other carbohydrates are more easily affected.
    Deacetylation (saponification)Deacetylation (also called saponification) reverses the effects of acetylation in some, if not all of the materials which were blocked. The reversal is progressive, that is, some substances recover their stainabilty before others.
    MethylationExtended methylation causes some otherwise oxidisable carbohydrates to become non-reactive. Primarily, however, it inhibits basophilia.
    Aldehyde blockIf applied before periodic acid oxidation, pre-existing aldehydes are inhibited from reacting and will be unstained. If applied after periodic acid oxidation, aldehydes produced by the periodic acid are inhibited from reacting and will be unstained. A second section which has not been blocked can be used to prove that oxidation formed the aldehydes.
    Amylase digestionAmylase (diastase) is used to remove glycogen. This is either to prove that a subtance is glycogen, or to remove it so that other PAS positive materials other than glycogen may be more clearly seen.
    Protease digestionProteases may also be used to remove carbohydrate-protein complexes, although this is less common than amylase digestion because the enzymes also digest other tissue proteins.
    Reducing rinsesThis is an acidified solution of potassium iodide and sodium thiosulphate applied following periodic acid to remove any iodate or periodate remaining in the tissue.
    Methanol, Pyridine & Solvent extractionPAS positive carbohydrate-lipid complexes, or glycolipids, may be removed with these solvents. They may be used at ambient or elevated temperature.
  4. Ensure sections are in water or an appropriate concentration of ethanol.
  5. Oxidize for the appropriate time in the selected periodic acid solution.
  6. Wash five minutes in running water, or 70% ethanol if using an alcoholic variant.
  7. Optionally, do one of the following.
    1. Hotchkiss’ reducing rinse, or
    2. Aldehyde block
  8. Place in Schiff’s reagent for 10 minutes, or longer if specially required.
  9. Transfer to three successive sulfite rinses, 2 minutes each.
  10. Wash in running tap water for 10 minutes.
  11. Optionally, stain nuclei.
  12. Optionally, stain cytoplasm.
  13. Dehydrate and differentiate with two changes each of 95% and absolute ethanol.
  14. Clear with xylene and coverslip using a resinous medium.

Expected Results

  • 1-2-glycols  –  red
  • Nuclei  –  blue or black
  • Cytoplasm  –  yellow or unstained

Notes

  • A strong Schiff’s reagent is recommended, i.e. one that contains 0.5 or 1 gram of pararosanilin per 200 mL solution.
  • Sections should be celloidinised before deacetylation, and it may be advisable after the other options in step 3.
  • Experience shows that the reducing rinses may be omitted.
  • Mayer’s hemalum or Weigert’s iron hematoxylin are suitable nuclear stains.
  • Sat. picric acid, 2% orange G, or picro-methyl blue are suitable cytoplasmic stains.

Safety Note

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

References

  1. Lillie, R.D., (1954)
    Histopathologic technique and practical histochemistry Ed.2
    Blakiston, New York, USA.
March 30, 2023
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Double Oxidation Schiff

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

Double Oxidation Schiff

14
steps
3
materials
print

Double oxidation may be used with any procedure which demonstrates aldehydes, including Schiff’s reagent and methenamine silver reduction. Its use is not necessarily confined to staining fungi and it may be found useful when demonstrating other structures and the background stains too darkly.

Materials

  • Periodic acid – 1% aqueous
  • Analine-acetic block
    MaterialAmount
    Acetic acid, glacial5mL
    Aniline oil45mL

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Oxidize with periodic acid for 10 minutes.
  3. Rinse well with tap water.
  4. Treat with aniline-acetic for 30 minutes.
  5. Rinse well with tap water.
  6. Re-oxidize with periodic acid for 20 minutes.
  7. Rinse well with tap water.
  8. Rinse with distilled water.
  9. Place in Schiff’s reagent for 10-30 minutes.
  10. Wash off with distilled water.
  11. Wash well with tap water for about 10 minutes.
  12. Counterstain with Mayer’s hemalum for 2 minutes.
  13. Wash well with tap water until hemalum is blued.
  14. Dehydrate with ethanol, clear with xylene and coverslip using a resinous medium.

Expected Results

  • Oxidizable carbohydrates  –  red
  • Fungi  –  red
  • Nuclei & background  –  blue

Notes

  • The time periodic acid is applied in the first oxidation determines the amount of background staining eliminated. If there is only a small amount of this material, then all of it may be blocked
  • The first oxidation will also oxidize some of the target material, and this staining will also be inhibited. If the first oxidation is applied for too long, the depth of staining of the target material may be affected. The 10 minutes specified is usually enough.
  • The time periodic acid is applied in the second oxidation determines the depth of staining for the target material. 20 minutes is usually adequate, but extending it may give darker staining. At some point, extending the time in periodic acid will cease to produce darker staining as all available carbohydrate will have been oxidized.

Safety Note

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

References

  1. Personal observation, B. Llewellyn,
March 30, 2023
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Bauer Reaction for Carbohydrates

By Aldehydes, Chromic Acid-Schiff Reaction, Protocols, Schiff's Reagent Reactions, Stain Target, Stain Type
Protocol

Bauer Reaction

for Carbohydrates

9
steps
7
materials
print

Materials

  • A Schiff reagent
  • A progressive hemalum, such as Mayer
  • Chromic acid
    MaterialAmount
    Chromium trioxide4g
    Distilled water100mL
  • Sulfurous acid
    MaterialAmount
    Sodium metabisulfite, 10% aqu.6mL
    Hydrochloric acid, 1N5mL
    Distilled water100mL

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Other fixatives are likely to be satisfactory. Most trichrome stains benefit from picric acid or mercuric chloride fixation. Formalin fixed tissues may benefit from secondary fixation of sections in Bouin’s fluid.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Oxidize in chromic acid for 40-60 minutes.
  3. Rinse with tap, then distilled water.
  4. Place into Schiff’s reagent for 15 minutes.
  5. Place into sulfurous acid rinses, 3 changes of 2 minutes each.
  6. Wash with running tap water.
  7. Counterstain with hemalum for 1 minute, and blue
  8. Dehydrate with ethanols.
  9. Clear with xylene and mount with a resinous medium.

Expected Results

  • Glycogen, mucin  –  red
  • Fungi  –  red
  • Nuclei  –  blue

Notes

  • Modern practice is to leave out the sulfite rinses and wash with large amounts of tap water.
  • A progressive hemalum should be used as counterstain because regressive hemalums sometimes stain mucin.
  • Mucins are not usually as dark as with a PAS.
  • Applying chromic acid for too long weakens staining due to continued oxidation of the aldehydes first produced.

Safety Note

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

References

  1. McManus, J. F. A. and Mowry, R. W., (1960)
    Staining Methods Histologic and Histochemical
    Harper & Row, New York, NY, USA.
March 30, 2023
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Jones’ Impregnation for Basement Membranes

By Aldehydes, Carbohydrates, Metal Impregnation, Metal Impregnation, Silver, Protocols, Stain Target, Stain Type
Protocol

Jones' Impregnation

for Basement Membranes

13
steps
9
materials
print

Materials

  • Periodic acid, 0.5% aqu.
  • Neutral red, 1% aqu.
  • Yellow gold chloride, 0.2% aqu.
  • Light green SFy, 0.2% in 0.2% acetic acid, or progressive hemalum and eosin
  • Sodium thiosulfate, 2.5% aqu.
  • Stock Methenamine silver
    MaterialAmount
    Methenamine, 3% aqu.100mL
    Silver nitrate, 5% aqu.5mL

    Shake until the precipitate redissolves. Silvering of the container indicates deterioration.

  • Working Methenamine silver
    MaterialAmount
    Stock Methenamine silver50mL
    Borax, 5% aqu.5mL

    Make just before use and preheat to 50°C.

Tissue Sample

3µ paraffin sections of neutral buffered formalin or Bouin fixed tissue are suitable. Other fixatives are likely to be satisfactory. A section adhesive is recommended. Thinner sections are to be preferred. This method gives excellent results with deplasticized methyl methacrylate sections at 1µ.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Oxidize with 0.5% periodic acid for 15 minutes.
  3. Rinse well with tap water.
  4. Rinse with distilled water.
  5. Treat with methenamine silver solution at 50&degC. until impregnated (up to 3 hours)
  6. Wash with distilled water.
  7. Tone with 0.2% gold chloride solution for 2 minutes.
  8. Rinse with distilled water.
  9. Fix in 2.5% sodium thiosulfate for 3 minutes.
  10. Wash well with running tap water.
  11. Counterstain with light green, neutral red or a light H&E.
  12. Rinse with tap water.
  13. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Basement membranes  –  black
  • Oxidisable carbohydrates  –  black
  • Background  –  as counterstained

Notes

  • In order to see the basement membranes on edge it is necessary to use the thinnest sections possible, especially for glomeruli.
  • Methenamine is also known as hexamethylenetetramine and hexamine.
  • Borax is sodium tetraborate.
  • Toning is a variable step. Untoned sections give dark brown material on a paler brown background. Many microscopists prefer to tone for about 15 seconds to produce brown-black material on a pale grey-brown background. Others tone longer (a few minutes) to produce black material on a grey background. Longer toning produces purple tones. Tone according to the personal preference of the microscopist reviewing the slides.
  • This method depends on a similar principle to the periodic acid Schiff reaction, but in which the aldehydes produced by oxidation reduce a silver solution instead of combining with Schiff’s reagent to form a red compound. Consequently, those materials which are red in a PAS will be black in Jones’ stain, i.e. it is not specific for basement membranes but will demonstrate any carbohydrates which can be oxidised to aldehydes.
  • Hayashi, Tome and Shimosato recommended that, after oxidation with periodic acid, thiosemicarbazide should be applied to the section. Thiosemicarbazide has the formula H2NNHCSNH2. The hydrazine group (H2NNH-) combines with any aldehydes generated by periodic acid oxidation. The thiocarbamyl group (-CSNH2) is a more powerful reducing agent than the aldehydes it replaces and reduces the methenamine silver solution more rapidly and with higher contrast.Immediately following step 3:
    • Place sections in 1% aqueous thiosemicarbazide for 10 minutes.
    • Wash well with tap water, and carry on from step 4.
  • It is well known that metallic azides can be explosive. However, thiosemicarbazide is not a simple metallic azide. The MSDS says:
    • Flash Point: n/a
    • Lower Explosive Limit: n/a
    • Upper Explosive Limit: n/a
    • Unusal Fire and Expl.rds: none identified

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. Hayashi, I., Tome, Y. and Shimosato, Y., (1989)
    Thiosemicarbazide used after periodic acid makes methenamine silver staining of renal glomerular basement membranes faster and cleaner.
    Stain Technology, v 64, p 185.
March 30, 2023
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Naphthoic Acid Hydrazide for Aldehydes

By Aldehydes, Protocols, Stain Target
Protocol

Naphthoic Acid Hydrazide

for Aldehydes

11
steps
11
materials
print

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.
March 30, 2023
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Fluorescent Schiff

By Aldehydes, Chromic Acid-Schiff Reaction, Protocols, Schiff's Reagent Reactions, Stain Target, Stain Type
Protocol

Fluorescent Schiff

11
steps
7
materials
print

This method demonstrates fungi, and may also be called a Fluorescent Gridley or CAFS.

Materials

  • Fluorescent Schiff reagent
  • Solution A
    MaterialAmount
    Chromium trioxide50g
    Distilled water500mL
  • Solution B
    MaterialAmount
    Sodium sulfite5g
    Distilled water500mL
  • Solution C
    MaterialAmount
    Ethanol, 70%495mL
    Hydrochloric acid5mL

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 solution A for 10 minutes.
  3. Wash well with tap water.
  4. Bleach with solution B for 1 minute.
  5. Wash well with tap water.
  6. Rinse with distilled water.
  7. Place in fluorescent Schiff reagent for 20 minutes.
  8. Wash well with tap water.
  9. Place in solution C, two changes, 5 minutes each.
  10. Wash well with tap water.
  11. Dehydrate, clear and mount in a non fluorescent resinous medium.

Expected Results

Using a BG 12 exciter filter, and OG 4 (yellow) and/or OG5 (orange) barrier filter, fungi fluoresce yellow with acriflavine and green-red with acridine orange.

Notes

  • This is most useful as a screening method.
  • If background fluorescence is too bright for fungi to be distinguished,it may be quenched with an alum hematoxylin for 1 minute,or 0.5% aqueous potassium permanganate for 1 minute.Quench immediately before the final dehydration step.This should be done with caution as it may reduce fungal fluorescence.

Safety Note

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

References

No specific reference is given for this method. A similar technique using periodic acid as the oxidant is found in:

  1. Culling C.F.A., (1974)
    Handbook of histopathological and histochemical techniques Ed. 3
    Butterworth, London, UK.
March 30, 2023
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Periodic Acid Fluorescent Schiff

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

Periodic Acid Fluorescent Schiff

9
steps
2
materials
print

Materials

Tissue Sample

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

Protocol

  1. Dewax and hydrate sections.
  2. Place in periodic acid for 10 minutes.
  3. Wash well with tap water.
  4. Rinse with distilled water.
  5. Place in fluorescent Schiff reagent for 20 minutes.
  6. Wash well with tap water.
  7. Place in acid alcohol, two changes, 5 minutes each.
  8. Wash well with tap water.
  9. Dehydrate, clear and mount in a non fluorescent resinous medium.

Expected Results

Using a BG 12 exciter filter, and OG 4 (yellow) and/or OG5 (orange) barrier filter, 1-2-glycols fluoresce yellow with acriflavine and green-red with acridine orange.

Periodic acid fluorescent schiff

Notes

  • The same materials fluoresce as would be red or pink in a regular PAS reaction.
  • If background fluorescence is too bright it may be quenched with alum hematoxylin for 1 minute,or 0.5% aqueous potassium permanganate for 1 minute. Quench immediately before the final dehydration step.This should be done with caution as it may reduce fluorescence.

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., (1974)
    Handbook of histopathological and histochemical techniques Ed. 3
    Butterworth, London, UK.
March 30, 2023
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