Skip to main content
Category

Protocols

[facetwp template="protocol_by_facet"]

Gram Weigert for Fibrin and Gram Positive Bacteria

By Bacteria, Fibrin, Gram Staining, Protocols, Stain Target, Stain Type

Gram Weigert

for Fibrin and Gram Positive Bacteria

11
steps
9
materials

Materials

Eosin

MaterialAmount
Eosin Y ws5g
Distilled water100mL

Crystal violet

MaterialAmount
Crystal violet1g
Distilled water100mL

Gram’s iodine

MaterialAmount
Iodine2g
Potassium iodide4g
Distilled water400mL

Aniline-Xylene

MaterialAmount
Aniline1volume
Xylene1volume

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 eosin solution for 5 minutes.
  3. Rinse with tap water.
  4. Place in crystal violet 1 minute.
  5. Rinse with tap water.
  6. Flood with Gram’s iodine for 1 minute.
  7. Rinse with tap water.
  8. Gently blot the section, being careful not to damage it.
  9. Decolorise the section with aniline-xylene.
  10. Rinse with several changes of xylene to remove all aniline.
  11. Mount with a resinous medium.

Expected Results

  • Gram positive bacteria  –  blue
  • Fibrin  –  blue
  • Background  –  pink

Notes

  • Control the differentiation with aniline-xylene microscopically. To examine, place the section in xylene to stop dye removal. Return to aniline-xylene if more differentiation is needed. Stop differentiation when the target element has good contrast.
  • Increasing the aniline content of the aniline-xylene will increase the speed of dye removal. Decreasing it will slow dye removal.
  • If the background is not pink enough, increase the time in eosin, stain in eosin at elevated temperature, or increase the eosin concentration.
  • The eosin counterstain may be omitted entirely if wished.

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., (1963)
    Handbook of histopathological techniques, 2nd ed.
    Butterworths, London.
  2. McManus, J.F.A. and Mowry, R.W., (1960),
    Staining methods, histologic and histochemical,
    Harper & Row, New York, NY, USA.

Baker’s Hematal Variants

By Dye Type, Hematoxylin and Eosin Staining, Mordanted Hematoxylin, Protocols, Stain Type

Baker's Hematal Variants

15
steps
11
materials

Materials

MaterialSolutionFunction
Stock AStock B
Aluminum sulphate15.76 gMordant
Distilled water1 LSolvent
Haematein1.876 gDye
Ethylene glycol, 50%, aqu.1 LSolvent
MaterialHematal-8Hematal-16
Stock A1 vol2 vol
Stock B1 vol1 vol
Ethylene glycol, 50%, aqu.1 vol
MaterialSolutionFunction
Standard AStandard B
Aluminum sulphate3.94 gMordant
Distilled water1 LSolvent
Haematoxylin5 gDye
Sodium iodate0.5 gDye
Distilled water1 LSolvent
Sulphuric acid, 0.5% aqueousas needed
Ammonia, 0.5% aqueousas needed

Compounding Procedures

Hematal-8 and Hematal-16 Solutions

  1. Prepare each stock solution separately.
  2. Combine as specified.
  3. The solution may be used immediately.

Standard Solution

  1. Prepare each solution separately.
  2. Standard solution B should be brought to a boil, then immediately cooled.

Protocol

Hematal-8 & Hematal-16 Solutions

  1. Bring sections to water with xylene and ethanol.
  2. Place into the staining solution for the specified time.
    • Hematal-8 for 2-5 minutes.
    • Hematal-16 for 10 minutes.
  3. Rinse with water and blue.
  4. Rinse well with water.
  5. Counterstain if desired.
  6. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Standard Solutions

  1. Bring sections to water with xylene and ethanol.
  2. Place into standard solution A for 1 hour.
  3. Rinse with water.
  4. Place into standard solution B for 20 minutes.
  5. Rinse with water.
  6. Place into 0.5% sulphuric acid for 40 seconds.
  7. Rinse with water.
  8. Place into 0.5% ammonia water for 5 seconds.
  9. Wash well with water.
  10. Optionally, counterstain with eosin.
  11. 70% ethanol for 30 seconds.
  12. 90% ethanol for 30 seconds.
  13. Absolute ethanol, 2 changes, 1 minute each.
  14. Xylene, 2 changes, 1 minute each.
  15. Mount with a resinous medium.

Expected Results

  • Nuclei  –  blue
  • Background  –  as counterstain or unstained

Notes

  • Hematal-8 and Hematal-16 may be used for routine staining. The solutions are quickly and conveniently prepared.
  • Hematal-8 may be used regressively by applying for 30 minutes, then lightly differentiating in acid. Dilute sulphuric acid was recommended (0.5% aqu.), but hydrochloric acid was usable.
  • Hematal-8 has 8 atoms of aluminum for each molecule of hematein, Hematal-16 has 16, hence the names.
  • Blueing is done with alkaline solutions such as hard tap water, Scott’s tap water substitute, 0.1% ammonia water, 1% aqueous sodium acetate, 0.5% aqueous lithium carbonate etc.
  • The standard method was used to give consistent staining with the usually expected appearance, but with the ability to vary each step for experimental reasons. It was not intended for routine use.

Safety Note

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

References

  1. Baker, J. R., (1962),
    Experiments on the action of mordants: 2. Aluminium-haematein.
    Quarterly Journal of Microscopical Science, v. 103, pt. 4, pp. 493-517.

Jones’ Impregnation for Basement Membranes

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

Jones' Impregnation

for Basement Membranes

13
steps
9
materials

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.

Smith’s Vanadium Hematoxylin

By Dye Type, Hematoxylin and Eosin Staining, Mordanted Hematoxylin, Protocols, Stain Type

Smith's Vanadium Hematoxylin

7
steps
5
materials

When differentiated with dilute lithium carbonate, this solution demonstrates basic proteins, including nuclear histones.

Materials

MaterialAmountFunction
Hematoxylin50 mgDye
Ethanol, 100%5 mLSolvent
Glycerol10 mLSolvent
Distilled water35 mLSolvent
Ammonium metavanadate200 mgSolvent

Compounding procedure

  1. Dissolve the hematoxylin in the ethanol, then add the glycerol and water in sequence.
  2. Add the ammonium vanadate and stir for 30 minutes.
  3. The ammonium vanadate may not completely dissolve. The pH is 6.2.

Tissue Sample

Paraffin sections of Schaudinn, Bouin or formalin fixed tissues are suitable. Sections of Zenker or Helly fixed tissues must be soaked in saturated aqueous lithium carbonate for 4 hours to stain satisfactorily.

Protocol

  1. Bring sections to water with xylene and ethanol.
  2. Place into the vanadate hematoxylin for 30 minutes.
  3. Place directly into 0.08% aqueous lithium carbonate for the appropriate time.
    Differentiation times in lithium carbonate

    FixativeTime
    Bouin2 minutes
    Formalin, 10%2 minutes
    Helly10 minutes
    Schaudinn5 minutes
    Zenker10 minutes
  4. Briefly rinse with distilled water for 2 seconds.
  5. Optionally, counterstain if wished.
  6. Dehydrate with 70%, 95% and absolute ethanols.
  7. Clear with xylene and mount with a resinous medium (Permount specified).

Expected Results

  • Nuclear histone  –  blue
  • Ribosomal protein  –  paler blue
  • Other protein  –  as counterstain or unstained

Notes

  • Counterstaining can be accomplished at step 5 with 1% aqueous eosin Y, phloxine B or erythrosin B.
  • Alternatively, metanil yellow at 0.1% in 95% ethanol containing 0.1% acetic acid may be used between the 70% and 95% ethanols in step 6.
  • Counterstaining with 0.01% aqueous safranin O stained RNA intensely, giving good contrast between blue nuclear histone and endoplasmic reticulum.

Safety Note

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

References

  1. Smith, A. A., (1995)
    A vanadate hematoxylin stain for basic proteins.
    Biotechnic and Histochemistry, v. 70, Nº 5, p. 5.

DMAB-Nitrite for Tryptophan

By Amyloid, Protocols, Stain Target

DMAB-Nitrite for Tryptophan

6
steps
2
materials

The DMAB-nitrite histochemical method is a simple and very highly selective, almost specific, method for the amino acid, tryptophan. A blue product obtained with this technique is invariably accepted as proof that the blue stained material contains tryptophan.

DMAB is p-dimethylaminobenzaldehyde, also known as 4-dimethylaminobenzaldehyde, 4-Formyl-N,N-dimethylaniline and N,N-Dimethyl-4-formylaniline.

Tryptophan

Tryptophan

DMAB

p-Dimethylaminobenzaldehyde

Beta Carboline

Product: Beta Carboline

The leftmost structural formula above is of tryptophan, the central one is of DMAB, and the rightmost formula is of the initial reaction product that is produced by them. This reaction product is then oxidized with potassium nitrite to form an easily seen, blue colored compound of unspecified structure. It should be noted that formaldehyde and glyceraldehyde can also participate in similar reactions, so if this method is contemplated then formalin fixation should be kept short. The common overnight fixation in 10% formalin variants still permits the technique to give a positive result.

Materials

  • p-Dimethylaminobenzaldehyde, 5% in concentrated hydrochloric acid
  • Sodium nitrite (NaNO2), 1% in concentrated hydrochloric acid

Tissue Sample

15µ paraffin sections of tissues fixed up to 24 hours in one of:

  • 1% trichloracetic acid in 80% ethanol
  • 10% sulphosalicylic acid
  • 10% neutral buffered formalin (6 hours preferred)

Protocol

  1. Bring sections to ethanol via xylene and ethanol and allow to just become dry.
  2. Place into DMAB solution for 1 minute
  3. Place in sodium nitrite solution for 1 minute.
  4. Rinse with tap water for 30 seconds.
  5. Dehydrate with ethanol.
  6. Clear with xylene and mount with a resinous medium.

Expected Results

  • Tryptophan  –  blue

Notes

  • Positive results should be seen in fibrin and fibrinoid, amyloid, Paneth cell granules, peptic cell granules, zymogen granules, muscle, neurokeratin and hair root sheath. Of these, only fibrin, including fibrinoid, and amyloid are extracellular materials which could be confused.
  • When used to confirm that a material is amyloid, the positive material may be differentiated from fibrin by dye staining methods, i.e. a positive DMAB-nitrite stain with a congo red stain also positive would identify the stained material as amyloid with a high degree of certainty, as fibrin is not stained with that dye.

Safety Note

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

References

  1. Adams, C.W.M., (1957),
    A p-dimethylaminobenzaldehyde-nitrite method for the histochemical demonstration of tryptophane and related compounds.,
    Journal of Clinical Pathology, v 10, page 56-62.
  2. Pearse, A.G.E. (1968).
    Histochemistry: Theoretical and Applied, Ed. 3, Volume 1.
    Churchill Livingstone, London, England.
  3. Davenport, H.A.. (1960).
    Histological and Histochemical Technics,
    W. B. Saunders, Philadelphia, USA.

HPS, AFIP Modification General Oversight stain

By Protocols, Stain Type, Trichrome Staining, Yellowsolve Staining

HPS, AFIP Modification

General Oversight stain

11
steps
6
materials

Also known as the Hematoxylin Phloxine Saffron stain, AFIP modification.

Materials

Tissue Sample

5 µ paraffin sections of neutral buffered formalin fixed tissue are suitable, as are many other fixatives.

Protocol

  1. Bring sections to water with xylene and ethanol.
  2. Place in picric acid solution for 5 minutes.
  3. Wash with water to remove yellow.
  4. Stain nuclei with hemalum, differentiate and blue.
  5. Wash well with water.
  6. Place in solution A for 2 minutes.
  7. Wash with tap water for 5 minutes.
  8. Thoroughly dehydrate with absolute ethanol.
  9. Place into solution B for 5 minutes.
  10. Rinse well with absolute ethanol.
  11. Clear in xylene and mount with a synthetic resinous medium.

Expected Results

  • Nuclei  –  blue
  • Muscle & cytoplasm  –  red
  • Collagen  –  yellow

Notes

  • The saffron should be extracted by mixing with anhydrous ethanol in a tightly capped container, then placing in a 56°C oven for a few days. Store and use at room temperature. It has a limited life and is best when freshly made.
  • 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.
  • It is important that there be no water in the saffron solution and that sections be thoroughly dehydrated before it is applied.
  • Usually, whole stigmata are more effective than ground saffron.
  • Erythrosine B may be used instead of phloxine B.

Safety Note

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

References

  1. Histological Staining Methods of the Armed Forces Institute of Pathology, 3rd ed. (1968)
    Luna, Lee G.
    McGraw-Hill, NY, USA

Akita & Kaneko’s Hemalum for Elastic Fibers

By Dye Type, Elastic Fibers, Hematoxylin and Eosin Staining, Mordanted Hematoxylin, Protocols, Stain Target, Stain Type

Akita & Kaneko's Hemalum

for Elastic Fibers

5
steps
6
materials

Materials

MaterialAmount
Hematoxylin100mg
Ethanol, 70%100mL
Sodium iodate20mg
Potassium alum300mg
Chloral hydrate5g
Citric acid100mg

Staining Solution Preparation

  1. Dissolve the hematoxylin into the ethanol.
  2. Add the potassium alum, then the other ingredients.
  3. Mix well for 15 to 20 minutes, then filter.
  4. At this point the solution is stable for 2 months at room temperature.
  5. Immediately prior to use, adjust to pH 8.0 with saturated aquueous lithium carbonate or 1N potassium hydroxide.

Tissue Sample

5µ paraffin sections of neutral buffered formalin fixed tissue are suitable. Helly and Bouin are also suitable. Other fixatives should be tested.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into freshly pH adjusted staining solution for 30-60 minutes.
  3. Wash with running tap water for 10 minutes.
  4. Dehydrate with ethanols.
  5. Clear with xylene and mount with a resinous medium.

Expected Results

  • Elastic fibres  –  blue
  • Collagen  –  pale violet

Notes

  • Note that this is an ethanolic variant of Mayer’s hemalum.
  • If sodium iodate is left out, the staining solution lasts 3-4 days, but the staining time is extended to 60 minutes.
  • The solution may also be made by diluting 30 mL Mayer’s hemalum with 70 mL absolute ethanol, mixing for 20 minutes, and filtering before adjusting the pH. Staining requires 60 minutes and the results are paler.

Safety Note

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

References

  1. Akita, M. and Kaneko, K., (1981)
    An improved hematoxylin method with application of Mayer’s hemalum for staining of elastic fibres.
    Stain Technology, v. 56, p. 327
  2. Kaneko, K. and Akita, M., (1978)
    On the staining of elastic fibres with alum hematoxylin.
    Stain Technology, v. 53, p. 43.

Lendrum, Slidders & Fraser’s Alcian Blue for amyloid

By Amyloid, Protocols, Stain Target

Lendrum, Slidders & Fraser's Alcian Blue

for amyloid

10
steps
10
materials

Materials

  • Stock A
    Alcian blue 8GX1g
    Ethanol 80%100mL
  • Stock B
    Sodium sulfate, hydrate1g
    Distilled water100mL
  • Working solution
    Stock A45mL
    Stock B45mL
    Glacial acetic acid10mL

    Mix. Leave 30 minutes. Prepare daily.

  • Acetic ethanol
    Ethanol, 95%45mL
    Distilled water45mL
    Glacial acetic acid10mL

    Prepare daily.

  • Borax Ethanol
    Boraxasrequired
    Ethanol, 80%100mL

    Dissolve borax to saturation.

Tissue Sample

Paraffin sections of formalin fixed tissues are satisfactory. If using the more complex trichrome counterstains, formal sublimate fixation is preferred.

Protocol

  1. Bring sections to water via xylene and ethanol.
  2. Place into acetic alcohol for 1 – 2 minutes.
  3. Place into the working solution for 2 hours.
  4. Place into acetic alcohol for 1 – 2 minutes.
  5. Wash with water.
  6. Place into alcoholic borax for 30 minutes.
  7. Wash with water.
  8. Counterstain (see notes).
  9. Dehydrate with absolute ethanol.
  10. Clear with xylene and mount with a resinous medium.

Expected Results

  • Amyloid  –  blue green
  • Other tissue  –  according to the counterstain

Notes

  • The simplest counterstain is the van Gieson.
  • If counterstaining with PAS, treatment with alcoholic borax may be omitted.
  • If counterstaining with silver impregnation for reticulin, the method must not include a Mallory bleach.
  • The authors suggested their own, more complex trichrome counterstain.

Safety Note

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

References

  1. Lendrum, A.C., Slidders, W. and Fraser, S., (1972),
    Renal hyalin: A study of amyloidosis and diabetic fibrinous vasculosis with new staining methods.,
    Journal of Clinical Pathology, v 25, page 373.

Gabe’s Aldehyde Fuchsin

By Aldehyde Fuchsin, Dye Type, Elastic Fibers, Protocols, Stain Target, Stain Type

Gabe's Aldehyde Fuchsin

6
steps
9
materials

Materials

Stock powder

MaterialAmount
Pararosanilin5g
Paraldehyde5mL
Hydrochloric acid10mL
Distilled water1L

Dye Preparation Procedures

  1. Add the dye to the water and boil.
  2. Cool to room temperature.
  3. Add paraldehyde and hydrochloric acid.
  4. Ripen at RT two days. Filter. Wash precipitate with 50mL distilled water. Dry the ppt and paper at 60°C.
  5. Store in a labelled container. Stable for about ten years.

Working solution

MaterialAmount
Stock powder0.25g
Ethanol, 70%200mL
Acetic acid, glacial2mL

Acid ethanol

MaterialAmount
Hydrochloric acid, conc.1mL
Ethanol, 95%200mL

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 the working solution for 5 minutes.
  3. Rinse well with running tap water.
  4. Rinse with acid ethanol for 20-30 seconds to remove excess dye.
  5. Counterstain if wished.
  6. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Elastic fibres  –  purple
  • Mast cells  –  purple
  • Pituitary βcells  –  purple
  • Sulphated mucins  –  purple
  • Background  –  as the counterstain
  • Nuclei  –  as the nuclear stain

Notes

  • The basic fuchsin used for this solution should be one that is suitable for Schiff’s reagent, i.e., it should have a high pararosanilin content. Both methods involve forming a compound between an aldehyde and dye.
  • Light counterstaining with a progressive alum hematoxylin and eosin is also suitable.
  • Many other counterstains can be used, including methods such as Masson’s trichrome.

Safety Note

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

References

  1. Kiernan. J.A., (1999)
    Histological and histochemical methods: Theory and practice, Ed. 3
    Butterworth Heinemann, Oxford, UK.
    Citing:
    Gabe, M., (1976)
    Histological techniques.
    Masson, Paris.

Gomori’s Aldehyde Fuchsin

By Aldehyde Fuchsin, Dye Type, Elastic Fibers, Intracytoplasmic Granules, Mast Cells, Protocols, Stain Target, Stain Type

Gomori's Aldehyde Fuchsin

7
steps
4
materials

Materials

Solution A

MaterialAmount
Basic fuchsin1g
Paraldehyde, fresh1mL
Hydrochloric acid, conc.1mL
Ethanol, 70%200mL

Compounding Procedure

  1. Dissolve the dye in the ethanol.
  2. Add paraldehyde and hydrochloric acid.
  3. Ripen at room temperature for 48-72 hours.
  4. Refrigerate. The solution is stable for 2-3 months.

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. Wash with water.
  3. Rinse with 70% ethanol.
  4. Place in the staining solution for 10 minutes.
  5. Rinse well with 95% ethanol.
  6. Counterstain the nuclei and/or the cytoplasm if wished.
  7. Dehydrate with ethanol, clear with xylene and mount with a resinous medium.

Expected Results

  • Elastic fibres  –  purple
  • Mast cells  –  purple
  • Pituitary β cells  –  purple
  • Sulphated mucins  –  purple
  • Background  –  as the counterstain
  • Nuclei  –  as the nuclear stain

Notes

  • The basic fuchsin used for this solution should be one that is suitable for Schiff’s reagent, i.e., it should have a high pararosanilin content. Both methods involve forming a compound between an aldehyde and dye.
  • Light counterstaining with a progressive alum hematoxylin and eosin is also suitable.
  • Many other counterstains can be used, including methods such as Masson’s trichrome.
  • Gabe described a technique for the preparation and use of aldehyde fuchsin powder.

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.