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HISTOLOGY

LECTURE # 29

INTRODUCTION TO SPECIAL STAINS TECHNIQUES: NERVE TISSUE STAINING Rationale: Special stain techniques are one of the major processes done in the histology laboratory. These techniques are performed to be evaluated with the diagnostics slides from H&E’s. These stains are used as an aid and a diagnostic tool for a final diagnosis.

Objective: Once completed this lecture, the student should be able to: a)

Describe the different types of nerve tissue staining.

b)

Learn the various methods of stain demonstration.

c)

Learn the classification for Neurofibrils, axon, glial, astrocytes and myelin sheath staining.

d)

Learn the procedures and diagnostic tools for each stain.

NERVE TISSUE Introduction •

Nerve Tissue Nerve tissue can be divided into two anatomical parts: – Central Nervous System (CNS) • Brain & Spinal cord – Peripheral Nervous System (PNS) • All other nervous system

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In histology, nervous tissue consists of cells and cells processes and the various components of nerve tissues. These normally fall in three groups: – Neuronal cells and processes – Glial cells and processes – Myelin sheath

Neurons 1. Nissl Substances - basophilic material in the cytoplasm of the neuron 2. Nerve Cell Processes – Dendrites & Axons

Neuroglia 1. Oligodendroglia – small cells that function in the CNS to produce & probably maintain, the myelin sheath surrounding many axons 2. Astrocytes – stellate cells of two types: Protoplasmic, which occurs in the gray matter, and fibrous which occurs in the white matter 3. Microglia – fixed phagocytic cells found throughout the brain and spinal cord 4. Ependymal cells – true epithelial cells that line the ventricles and spinal canal, forms a barrier between the cerebrospinal fluid and nervous tissue

Myelin 1. Is a complex, white, fatty, nonliving material containing protein, cholesterol, phospholipids, and cerebrosides. 2. It is largely loss during routine paraffin processing with only neurokeratin, a resistant proteolipid, left in the embedded block. 3. It is formed by oligodendroglia in the central nervous system and by Scwann cells in the peripheral nervous system. These are normally demonstrated by Luxol Fast Blue and Iron hematoxylin

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Nissl Substance: Cresyl Echt Violet Method (Luna, 1960) I. Purpose of the stain: Identification of neurons in tissue sections, or demonstration of loss of Nissl substances (chromatolysis). II. Principle of the stain: They are very basophilic and will stain sharply with basic aniline dyes. By varying the pH either Nissl substances can be stained by itself or nuclei and Nissl substances can be stained. III. Fixatives: 10% Neutral buffered formalin is preferred. IV. Sectioning: Paraffin sections at 6-8 microns. V. Controls: Section of spinal cord VI. Reagents: 1. Cresyl Echt Violet Solution: Cresyl echt violet, distilled water. Need to be ripen for 24 to 48 hors and filtered before use. 2. Balsam-Xylene Mixture: Canada Balsam & Xylene VII. Procedure: 1. Deparaffinize and hydrate slides to water. 2. Stain slides for 3-5 minutes in Cresyl Echt Violet solution. This is the step for the staining of the nissl substances. 3. Rinse slides in two changes of distilled water. 4. Place sections in 95% alcohol for 30 seconds. 5. Place sections in absolute alcohol for 30 seconds. 6. Place section in xylene for 1 minute. 7. Place sections in balsam-xylene solution for 2 minutes. 8. Differentiate slides in two changes of absolute alcohol for 10 to 30 seconds each and check microscopically. 9. Take through several changes of xylene. Steps 7 – 9 may have to be repeated constantly. 10. Coverslip with synthetic resin media. VIII. Results: Nissl Substances..………………………………………….. Blue to purple

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Nissl Substance: Cresyl Echt Violet Method (Vacca) I. Purpose of the stain: Identification of neurons in tissue sections, or demonstration of loss of Nissl substances (chromatolysis). II. Principle of the stain: This procedure uses cresyl echt violet at an acid pH, restricting staining to DNA- and RNA-containing structures. III. Fixatives: 10% Neutral buffered formalin is preferred. IV. Sectioning: Paraffin sections at 6-8 microns. V. Controls: Section of spinal cord VI. Reagents: 1. Cresyl Echt Violet Solution: Cresyl echt violet, distilled water. Need to be ripen for 24 to 48 hors and filtered before use. 2. Balsam-Xylene Mixture: Canada Balsam & Xylene

VII. Procedure: 1. Deparaffinize and hydrate slides to water. 2. Stain slides for 8 minutes in working Cresyl Echt Violet solution. This is the step for the staining of the nissl substances. 3. Dehydrate sections in two changes of 95% alcohol & absolute alcohol. 4. Clear in two changes of xylene & coverslip. Results:

Nissl Substances & Nuclei ..………………………………………….. Blue purple

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Nerve fibers, Nerve endings, and Neurofibrils Bodian Method I. Purpose of the stain: Staining of nerve fiber in tissue sections. II. Principle of the stain: This procedure uses a silver proteinate known as Protargol™, which is mixed in with several shots of copper creating a chemical reaction suitable to the staining of the nerve fibers. Hydroquinone is the reducing reagent in this procedure. III. Fixatives: 10% Neutral Buffered Formalin is preferred IV. Sectioning: Cut Paraffin sections from 6 to 8 microns. V. Controls: Best control is peripheral nerve or cerebral cortex. Spinal cord is not a good control, since it will appear in a cross section. VI. Reagents: 1. 1% Protargol™: Protargol™ & Distilled water. 2. Reducing solution: hydroquinone, Formaldehyde & distilled water. 3. 1% Gold chloride solution: gold chloride & distilled water. 4. 5% Sodium thiosulfate (Hypo): sodium hyposulfite & distilled water. 5. Aqua Regia: Hydrochloric acid concentrated, nitric acid concentrated. Prepare and use under a hood. 6. Aniline blue solution: Aniline blue, oxalic acid, Phosphomolybdic acid & distilled water. VII. Procedure: 1. Deparaffinize and hydrate sections to distilled water. 2. For every 100 mL of Protargol solution, add 4 to 6g of clean copper shot (cleaned with aqua regia and rinsed well with distilled water). Place slides in this solution and let stand at 37°C for 48 hours. 3. Rinse sections in three changes of distilled water. 4. Place slides in the reducing solution for 10 minutes. 5. Rinse in three changes of distilled water. 6. Tone sections in gold chloride solution for 10 minutes. This solution may be reused. 7. Rinse in three changes of distilled water. 8. Develop in oxalic acid solution, checking with the microscope, until the background is gray and the nerve fibers appear clearly stained (approximately 3 to 5 minutes). Oxalic acid treatment should not be prolonged since over treatment will ruin the silver proteinate reaction. 9. Rinse in three changes of distilled water. 10. Treat sections with sodium thiosulfate for 5 minutes. 11. Rinse in distilled water. 12. Counterstain, if desired, with aniline blue solution (2 or 3 quick dips to give a light blue background). See note 4. 13. Dehydrate in 95% alcohol and absolute alcohol, two changes each. 14. Clear in two changes of xylene. 15. Mount with synthetic resin.

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VIII. Results: Nuclei……………………………..Black Background.....................................Light Gray or Blue Nerve Fibers................................... Black Bodian Stain Nerve Fibers

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Holmes Silver Nitrate Method I.

Purpose of the stain: The demonstration of nerve fibers and Neurofibrils in tissue section.

II.

Principle of the stain: Holmes attributed inconsistent results obtained with the Bodian technique to the fact that the Protargol solution never reaches the alkalinity necessary for optimal impregnation, and he modified the technique by developing a buffered impregnating solution. The pyridine in the solution is an alkali, and Holmes thought that this modified the electrostatic condition of the tissue. This is an argyrophil silver method, requiring that chemical reduction be used. The purposes of gold chloride, oxalic acid, and sodium thiosulfate are identified in the description of the Bodian procedure.

III.

Fixatives: 10% Neutral Buffered Formalin is preferred.

IV.

Sectioning: Cut Paraffin sections from 10 to 15 microns.

V.

Controls: Use a section of cerebral cortex. Spinal cord is not a good control for this stain as all axon appear in cross section.

VI.

Reagents: a) Aqueous Silver Nitrate, 20%: Silver nitrate & Distilled water b) Aqueous Silver Nitrate, 1 %: Silver nitrate: 20% Silver solution & Distilled water c) Boric Acid Solution: Boric acid & Distilled water d) Borax Solution: Sodium borate & distilled water e) 10% Pyridine Solution: Pyridine & Distilled water f) Impregnating Solution 1. Boric acid solution (fresh) 2. Borax solution (fresh) 3. Distilled water 4. Silver nitrate, 1 % aqueous solution 5. Pyridine, 10% aqueous solution Mix boric acid solution and borax solution in a 500-mL flask. Add the water, then the aqueous silver nitrate, and then the aqueous solution of pyridine. Mix thoroughly. Make enough solution for 20 mL per slide. g) Reducing Solution: Make fresh before use 1. Hydroquinone 2. Sodium sulfite (crystals) 3. Distilled water h) Gold Chloride, 0.2%: Gold chloride, 1 % solution & Distilled water i) Oxalic Acid Solution, 2%: Oxalic acid & Distilled water j) Sodium Thiosulfate (Hypo): Sodium thiosulfate & Distilled water

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VII. Procedure: 1. Deparaffinize sections and hydrate to distilled water. 2. Place sections in 20% silver nitrate in the dark at room temperature for 1 hour. 3. Prepare the impregnating solution. 4. Take slides from 20% silver nitrate and wash for 10minutes in three changes of distilled water. 5. Place slides in impregnating solution, allowing at least 20 mL of solution per slide. Cover jar and incubate overnight at 37°C. 6. Remove slides, shake off superfluous fluid, and place in the reducer for not less than 2 minutes. 7. Wash in running water for 3 minutes. 8. Rinse slides in distilled water. 9. Tone in 0.2% aqueous gold chloride for 3 minutes. This solution may be reused until a brown precipitate forms or the solution becomes cloudy. 10. Rinse in distilled water. 11. Place slides in 2% aqueous oxalic acid for 3 to 10 minutes. When the axons are thoroughly blue-black, stop the process. 12. Rinse in distilled water. 13. Place the slides in 5% aqueous sodium thiosulfate. 14. Wash in tap water for 10 minutes. As with Bodian stain, a counterstain may be applied at this point. 15. Dehydrate in two changes each of 95% alcohol and absolute alcohol. 16. Clear in xylene and mount with synthetic resin.

VIII. Results: Axon & Nerve Fibers……………………..Black Neurofibrils.................................................Black

Holmes Silver Nitrate

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Nerve Fibers, Neurofibrillary Tangles, and Senile Plaques: Bielschowsky Method I. Purpose of the stain: To demonstrate Nerve Fibers and the presence of neurofibrillary tangles and senile plaques in Alzheimer’s disease. II. Principle of the stain: The tissue is impregnated with the Ammoniacal silver solution. The silver deposited on Neurofibrils and axons is then reduced to metallic silver by the formaldehyde in the developer. Since the sections are not toned with gold chloride in this procedure, the yellow background remains, Sodium thiosulfate removes and unreduced silver. III. Fixatives: 10% Neutral Buffered Formalin. IV. Sectioning: Cut Paraffin sections from 8 microns. V. Controls: Tissue from the central nervous system must be used. If possible, the tissue should contain senile plaques and neurofibrillary tangles. VI. Reagents: 1. 1% Silver Nitrate: Silver nitrate & Distilled water: Prepare fresh. 2. 5% Silver Nitrate: Silver nitrate & Distilled water: Store in refrigerator. 3. 10%Nitric acid solution: Nitric acid & distilled water: Prepare fresh. 4. Developer: a) Formaldehyde: 37 to 40% b) Distilled water c) Citric Acid d) 10% Nitric acid Prepare Fresh 5. 1% Ammonium hydroxide solution: Ammonium hydroxide, Conc. & Distilled water. 6. 2% Sodium thiosulfate (Hypo): sodium hyposulfite & distilled water. VII. Procedure: 1. Deparaffinize the slides and hydrate to distilled water. 2. Place slides in 40 mL of 1.0% silver nitrate solution in a plastic Coplin jar and microwave at power level 3 (180 W) for 1 minute. Dip the slides up and down several times and allow them to remain in the warm solution (50°C) for 15 minutes. 3. Place the slides in distilled water. 4. Pour the warm 1 % silver nitrate used in step 2 into a 125-mL flask. Add 28% ammonium hydroxide drop by drop with constant shaking, until the initial precipitate disappears and the solution turns clear. Then add 5% silver nitrate drop by drop with constant shaking, until the solution becomes slightly cloudy. 5. Pour the ammoniacal silver solution prepared in step 4 into a plastic Coplin jar. Place slides in this solution and microwave at power level 3 (180 W) for 1 minute. Dip the slides up and down several times and allow them to remain in the warm solution (60°C) for 15 minutes. 6. Place slides in 1 % ammonium hydroxide solution for not more than 20 seconds. 7. Add three drops of developer to the ammoniacal silver solution used in step 5.Quickly mix with a glass rod and immediately place the slides in the solution for about 3 minutes or until the tissue sections turn brown. The solution will turn a grayish color and a mirror of silver will form on the sides of the Coplin jar and sometimes on the slides, but not on the tissue sections. 8. Place slides in 1 % ammonium hydroxide solution for not more than 15 seconds. 9. Rinse in three changes of distilled water. 9

10. Wipe off the mirror of silver from both sides of the slides, taking care not to damage the tissue sections. 11. Place the slides in 2% sodium thiosulfate for 30 seconds. 12. Rinse slides in four changes of distilled water. 13. Dehydrate in 95% alcohol and absolute alcohol, two changes each. 14. Clear in two changes of xylene and mount with synthetic resin. VIII. Results: Axon…………………………………………...……………………..Brown to black Cytoplasmic Neurofibrils…………………………………………… Brown to black Neurofibrillary tangles and plaques of Alzheimer’s disease......…….Dark brown to black Neuromelanin………………………………………………………. Black Lipofuchsin…………………………………………………………. Brown to black

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Bielschowsky-PAS Method I. Purpose of the stain: To demonstrate the presence of neurofibrillary tangles and senile plaques in Alzheimer’s disease. II. Principle of the stain: The tissue is impregnated with the Ammoniacal silver solution, and silver is deposited on neurofibrils and axons. The silver is then reduced to metallic silver by the formaldehyde in the developer. Gold chloride is used to tone the section, and this step eliminates the yellow background. Sodium thiosulfate removes any unreduced silver. The Schiff’s reaction is used to stain both basement membranes and Amyloid in the plaques. III. Fixatives: 10% Neutral Buffered Formalin. IV. Sectioning: Cut Paraffin sections from 8 to 10 microns. V. Controls: Tissue from the central nervous system must be used. If possible, the tissue should contain senile plaques and neurofibrillary tangles. VI. Reagents: 1. 20% Silver Nitrate: Silver nitrate & Distilled water: Prepare fresh. 2. Ammoniacal Silver: 20% silver nitrate & Ammonium hydroxide drop by drop. 3. Developer: a. Formaldehyde: 37 to 40% b. Distilled water c. Citric Acid d. Nitric acid, Conc. 4. 0.5% Gold Chloride solution: Gold chloride & Distilled water. 5. 5% Sodium thiosulfate (Hypo): sodium hyposulfite & distilled water. 6. 1% Periodic Acid: Periodic acid & Distilled water. 7. Schiff’s Reagent VII. Procedure: 1. Prepare the Ammoniacal silver solution before beginning the procedure. 2. Deparaffinize the slides and hydrate to distilled water. 3. Place slides in 20% silver nitrate in the dark at room temperature for 20 minutes. 4. Remove slides from the silver nitrate and wash once in distilled water. 5. Place slides in the Ammoniacal silver solution at room temperature for 20 minutes. 6. Wash slides in ammonia water (4 drops concentrated ammonium hydroxide to 100 ml of distilled water). 7. While the slides are in the ammonia water, add 2 drops of developer to the Ammoniacal silver solution used in step 5 and mix well. 8. Place slides in the mixed developer-ammoniacal silver solution. The tissue should turn brown, average time is 3 minutes. 9. Wash well in ammonia water, then in distilled water. 10. Tone slides in gold chloride until the first gray appears, approximately 30 seconds. 11. Wash slides in ammonia water, then in distilled water for 1 minute. 12. Place slides in 5% sodium thiosulfate (hypo) for 30 seconds. 13. Wash slides in running tap water for 5 minutes. 14. Rinse sections well in distilled water. 15. Place slides in 1% periodic acid solution for 5 minutes. 16. Rinse slides well in two changes of distilled water. 17. Place sections in the Schiff’s reagent for 5 minutes. 18. Dehydrate slides in two changes of 95% alcohol and two changes absolute alcohol. 11

19. Clear in two changes of xylene and mount with synthetic resin. VIII. Results: Axon…………………………………………...……………………..Black Amyloid(plaque cores and vascular)…………………………………Magenta Neurofibrillary tangles and peripheral neurites or neuritic plaques.….Dark black Lipofuchsin…………………………………………………….……. Magenta

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Sevier-Munger Method I. Purpose of the stain: To demonstrate nerve fibers and the presence of neurofibrillary tangles and senile plaques in Alzheimer’s disease. II. Principle of the stain: The tissue is impregnated with the Ammoniacal silver solution. The silver is deposited on neurofibrils and axons and then is reduced to metallic silver by formaldehyde. Since the sections are not toned by gold chloride in this procedure, the yellow background remains. Sodium thiosulfate removes any unreduced silver. III. Fixatives: 10% Neutral Buffered Formalin. IV. Sectioning: Cut Paraffin sections from 6 to 8 microns. V. Controls: Tissue from the central nervous system must be used. VI. Reagents: 1. 20% Silver Nitrate: Silver nitrate & Distilled water. 2. 10% Silver nitrate: Silver nitrate & Distilled water. 3. Formalin Solution: 37-40% Formaldehyde & Distilled water. 4. Sodium carbonate solution: sodium carbonate & Distilled water. 5. Ammoniacal Silver: 20% silver nitrate & Ammonium hydroxide drop by drop. 6. 5% Sodium thiosulfate (Hypo): sodium hyposulfite & distilled water. VII. Reagents: 1. Deparaffinize the slides and hydrate to distilled water. 2. Preheat the 20% silver nitrate to 60oC for 15 minutes. Add the slides to the warm silver solution and let them remain in the oven for 15 minutes. 3. Rinse one slide at a time in distilled water and place in a clean, dry staining jar. 4. While shaking gently, add 10 drops of the formalin solution to the working Ammoniacal silver solution. Quickly pour this solution over the slides and let develop for 5 to 30 minutes until golden brown. Check microscopically for completeness of the reaction. Do not wash. Keep in motion during development to avoid precipitation. 5. Rinse slides well in three changes of fresh tap water. 6. Place in sodium thiosulfate solution for 2 minutes, and wash well in tap water. 7. Dehydrate, clear, and mount with synthetic resin. VIII. Results: Nerve endings…………………………………………...……………………..Black Neurofibrillary tangles and peripheral neurites or neuritic plaques.………......Black

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Glial Fibers Mallory PTAH Stain I. Purpose of the stain: The demonstration of glial fibers. II. Principle of the stain: The amount of phosphotungstic acid is far greater than the amount of hematein (20:1) in the staining solution, and it is believed that the tungsten binds all available hematein to give a blue lake. This lake provides the blue color to selected tissue components (glial fibers, nuclei, and to a certain extent, myelin). The red-brown- or salmon-colored components (neurons) are believed to be stained by the phosphotungstic acid. Components will lose their redbrown color after water or prolonged alcohol washing, so the dehydration steps following staining should be rapid. III. Fixatives: 10% Neutral Buffered Formalin. IV. Sectioning: Cut Paraffin sections from 6 to 8 microns. V. Controls: Use a section of cerebral cortex (not spinal cord) for the demonstration of glial fibers. VI. Reagents: 1. PTAH Solution a) Hematoxylin b) Phosphotungstic acid c) Distilled water 2. Lugol Iodine: Iodine & distilled water Place the potassium iodide and about 150 mL of the water in a flask and stir until dissolved. Dissolve the iodine in this concentrated solution of potassium iodide. When the iodine is dissolved, add the remaining water and mix well. 3. 5% Sodium thiosulfate (Hypo): sodium hyposulfite & distilled water. 4. 1% Potassium Permanganate: Potassium permanganate & Distilled water. 5. 5% Oxalic Acid Solution: Oxalic acid & Distilled water. VII. Reagents: 1. Deparaffinize the sections and hydrate to distilled water. 2. Mordant the sections overnight at room temperature in Zenker solution containing acetic acid. 3. Wash the sections in running water for 15 minutes. 4. Place in Lugol iodine for 15 minutes. Do not take the slides through sodium thiosulfate (hypo) as this may impair the subsequent staining reaction. 5. Decolorize the sections in 95% alcohol for at least 1 hour. 6. Rinse rapidly in three changes of distilled water. 7. Place sections in 1 % potassium permanganate for 5 minutes. 8. Wash in running tap water for 10 minutes. 9. Decolorize the sections in 5% oxalic acid for 5 minutes. 10. Wash in running tap water for 10 minutes. 11. Stain in PTAH solution overnight at room temperature. 12. Dehydrate rapidly through two changes each of 95% and absolute alcohols, clear in xylene, and mount with synthetic resin.

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VIII. Results: Glial fibers….…………………………………………...……………………..Blue Nuclei………………………………………………………………………….Blue Neurons………………………………………………………………………..Salmon Myelin…………………………………………………………………………Blue

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Holzer Method I. Purpose of the stain: The demonstration of glial fibers and areas of gliosis. II. Principle of the stain: Glial fibers are stained with crystal violet and are resistant to decolorization with the alkaline aniline-chloroform mixture. III. Fixatives: 10% Neutral Buffered Formalin. IV. Sectioning: Cut Paraffin sections from 6 to 8 microns. V. Controls: Use a section of cerebral cortex (not spinal cord) for the demonstration of glial fibers. VI. Reagents: 1. Aqueous Phosphomolybdic Acid Solution: Phosphomolybdic acid & Distilled water. Prepare fresh. 2. Phosphomolybdic alcohol solution: .5% Phosphomolybdic acid & 95% alcohol. 3. Absolute alcohol-Chloroform Mixture: Absolute alcohol & Chloroform. Prepare fresh. a. Crystal Violet Stain, Prepare fresh: b. Crystal violet c. Absolute alcohol 4. Chloroform 5. Potassium bromide solution: potassium bromide & Distilled water. 6. Differentiating solution, Prepare fresh: a. Aniline oil b. Chloroform c. Ammonium hydroxide, concentrated VII. Procedure: 1. Deparaffinize the slides and hydrate to distilled water. 2. Place the sections in fresh phosphomolybdic acid-alcohol for 3 minutes. 3. Drain off the excess fluid, place slides on a staining rack, and cover the sections with absolute alcohol-chloroform mixture. The tissue should become translucent. 4. While the sections are still wet, cover them with crystal violet stain and allow to remain for 30 seconds. 5. Replace the stain with 10% Potassium bromide, washing for 1 minute with this solution. 6. Blot the section dry and then allow them to air-dry thoroughly. 7. Differentiate slides individually for 30 seconds in the differentiating solution. 8. Wash in several changes of xylene. Steps 7 & 8 may have to repeated several times until the background is very pale blue or colorless. 9. Mount with synthetic resin. VIII. Results: Glial fibers….……...…..Blue Background……………Very pale blue to colorless

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Astrocytes: Cajal Stain I. Purpose of the stain: The demonstration of astrocytes. This method has been replaced to a great extent by immunohistochemical procedures. II. Principle of the stain: Astrocytes are selectively stained with the Cajal gold sublimate method on frozen sections. III. Fixatives: Formalin ammonium bromide for no less than 2 days and no more than 25 days. If the tissue has been fixed originally in 10% neutral buffered formalin, wash and place in formalin ammonium bromide for 48 hours before proceeding with the technique. IV. Sectioning: Cut frozen sections at 20-30 microns. Do not pick up on slides; the section should be free-floating for this technique. Tissue will section better if washed in tap water for 30 minutes before freezing. V. Controls: Use a section of cerebral cortex (not spinal cord) for the demonstration of astrocytes. VI. Reagents: 1. Formalin Ammonium Bromide: Ammonium bromide, Formaldehyde 37-40% & Distilled water. 2. Gold Sublimate: 1% Gold Chloride, 1% Mercuric chloride & Distilled water. 3. 5% Sodium Thiosulfate (Hypo): Sodium thiosulfate & Distilled water. VII. Procedure: 1. Wash the free-floating frozen section in several changes of distilled water. 2. Transfer the sections to gold sublimate solution, and leave in the dark for 4 hours. The sections should be purple. 3. Wash well in distilled water. 4. Treat with 5% Sodium thiosulfate for 2 minutes. 5. Wash section well in several changes of distilled water. 6. Carefully mount the sections on slides, blot with bibulous paper, and dehydrate in 95% & 100% alcohols. 7. Clear in xylene and mount in synthetic resin. VIII. Results: Astrocytes with perivascular feet….…………………………………………………..Black

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Myelin Sheath: Weil Method I. Purpose of the stain: The demonstration of myelin in tissue. When an axon degenerates, the myelin sheath breaks down into simpler lipids; these simple lipids will be removed eventually. II. Principle of the stain: The mordant-hematoxylin solution attaches to the phospholipid component of the myelin sheath, which has an affinity for the cationic dye lake. This is a regressive staining technique with differentiation usually accomplished in two steps. The first differentiation is accomplished macroscopically with ferric ammonium sulfate (excess mordant), which removes most of the excess dye. The second differentiation is done microscopically with borax ferricyanide (oxidizer), which removes any remaining nonspecifically bound hematoxylin lake and forms a colorless oxidation product. Only the myelin sheath and red blood cells are left stained. III. Fixatives: 10% Neutral buffered formalin. IV. Sectioning: Cut paraffin section between 15 to 30 microns. V. Controls: Use a section of spinal cord or medulla. VI. Reagents: 1. Ferric Ammonium Sulfate Solution, 4%: Ferric ammonium sulfate & Distilled water 2. Alcoholic Hematoxylin, 10%: Hematoxylin powder & Absolute alcohol. 3. This solution should be allowed to stand for 2 to 3 days, but prolonged ripening is unnecessary, because the iron used in the staining solution is a strong oxidizer. 4. Staining Solution: Hematoxylin, 10% alcohol solution, distilled water. 5. Mix in an Erlenmeyer flask and add: Ferric ammonium sulfate, 4% solution. Prepare fresh. 6. Differentiating Solution: Sodium borate, Potassium ferricyanide & Distilled water. VII. Procedure: 1. Deparaffinize sections and hydrate to distilled water. 2. Transfer sections to the staining solution and stain for 30 minutes at 54°C to 56°C. 3. Wash in two changes of tap water. 4. Differentiate in 4% ferric ammonium sulfate until the gray matter can just be distinguished from the white matter and the stain is removed from the slides. 5. Wash in three changes of tap water. 6. Complete differentiation of the sections in sodium borate-potassium ferricyanide solution. This differentiation should be controlled microscopically until the gray and white matters are sharply defined. 7. Wash sections in two changes of tap water. 8. Treat sections with diluted ammonia water (about 6 drops to 100 mL of water). 9. Wash in distilled water. 10. Dehydrate in two changes each of 95% and absolute alcohols. 11. Clear in xylene and mount with synthetic resin.

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VIII. Results: Myelin sheath. . . . . . . . . . . . . . . . . Blue to blue-black Background. . . . . . . . . . . . . . . . . . Light tan

Myelin Sheath: Luxol Fast Blue 1. Purpose of the stain: The demonstration of myelin in tissue. When an axon degenerates, the myelin sheath breaks down into simpler lipids; these simple lipids will be removed eventually. 2. Principle of the stain: Luxol fast blue, like alcian blue, is of the sulfonated copper phthalocyanine type, but is alcohol soluble, whereas alcian blue is water-soluble. Staining is due to lipoproteins, and the mechanism is one of an acid-base reaction with salt formation; the base of the lipoprotein replaces the base of the dye. 3. Fixatives: 10% Neutral buffered formalin. 4. Sectioning: Cut paraffin section between 10 to 15 microns. 5. Controls: A section of spinal cord or medulla provides a good control. 6. Reagents: a) 0.1% Luxol fast blue: Luxol fast blue MBSN, 95% alcohol: Dissolve the dye in alcohol, then add: 10% Acetic acid. b) 0.05% Lithium carbonate: Lithium carbonate& Distilled water. c) 70% Alcohol solution: Absolute alcohol & Distilled water. 7. Procedure: 1. Deparaffinize sections and hydrate to distilled water. 2. Place slides in Luxol fast blue solution and leave overnight at 56°C to 58°C. The container should be tightly capped as this alcoholic solution will evaporate readily. 3. Rinse sections in 95% alcohol to remove excess stain. 4. Rinse in distilled water. 5. Begin the differentiation by immersing the slides in lithium carbonate solution from 10 to 20 seconds. 6. Continue the differentiation in 70% alcohol solution until the greenish blue of the white matter 19

contrast sharply with the colorless gray matter. 7. Wash sections in distilled water. 8. Dehydrate in two changes each of 95% and absolute alcohols. 9. Clear in xylene and mount with synthetic resin. 8. Results: Myelin sheath. . . . . . . . . . . . ……………………………………………………….. . . . .Blue Background. . . . . . . ……………………………………………………….. . . . . . . . . . . Colorless

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Myelin Sheath and Nissl Substances Combined Stain: Luxol Fast Blue-Cresyl Echt Violet I. Purpose of the stain: The demonstration of both myelin and Nissl substances in tissue sections. Nissl substances are lost after cell injury, and if the axon degenerates, the myelin covering also breaks down. II. Principle of the stain: Luxol fast blue, like alcian blue, is of the sulfonated copper phthalocyanine type, but is alcohol soluble, whereas alcian blue is water-soluble. Staining is due to lipoproteins, and the mechanism is one of an acid-base reaction with salt formation; the base of the lipoprotein replaces the base of the dye. They are very basophilic and will stain sharply with basic aniline dyes. By varying the pH either Nissl substances can be stained by itself or nuclei and Nissl substances can be stained. III. Fixatives: 10% Neutral buffered formalin. IV. Sectioning: Cut paraffin section between 10 to 15 microns. V. Controls: A section of spinal cord or medulla provides a good control. VI. Reagents: a) 0.1% Luxol fast blue: Luxol fast blue MBSN, 95% alcohol: Dissolve the dye in alcohol, then add: 10% Acetic acid. b) 0.05% Lithium carbonate: Lithium carbonate& Distilled water. c) 70% Alcohol solution: Absolute alcohol & Distilled water. d) 10% Acetic acid: Acetic acid & Distilled water. e) Cresyl Echt Violet: Cresyl echt violet & Distilled water. Just before use add 15 drops of 10% Acetic acid, filter and pre-heat. This solution is not very stable. VII. Procedure: 1. Deparaffinize sections and hydrate to distilled water. 2. Place slides in Luxol fast blue solution and leave overnight at 56°C to 58°C. The container should be tightly capped as this alcoholic solution will evaporate readily. 3. Rinse sections in 95% alcohol to remove excess stain. 4. Rinse in distilled water. 5. Begin the differentiation by immersing the slides in lithium carbonate solution from 10 to 20 seconds. 6. Continue the differentiation in 70% alcohol solution until gray matter and white matter can be distinguished. Do not overdifferentiate. 7. Wash sections in distilled water. 8. Finish differentiating the sections by rinsing them briefly in lithium carbonate solution, then putting them in 70% alcohol. The greenish blue of the white matter contrast sharply with the colorless gray matter. 9. Rinse slides thoroughly in distilled water. 10. Place slides in Cresyl echt violet solution for 6 minutes. Filter and preheat cresyl echt violet solution to 57°C just before use. Keep hot during staining. 11. Differentiate in several changes of 95% alcohol. 12. Dehydrate in two changes each of 95% and absolute alcohols. Clear in xylene and mount with synthetic resin. VIII. Results: Myelin sheath. . . . . . . . . . . . ……………………………………………………….. . . . .Blue Nissl Substances…………………………………………………………………………..Violet Nuclei……………………………………………………………………………………...Violet 21

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Myelin Sheath and Nerve Fibers Combined Stain: Luxol Fast Blue-Holmes Silver Nitrate Method I. Purpose of the stain: The demonstration of both myelin and nerve fibers in the same tissue section. If the axon degenerates, the myelin covering also breaks down into simpler lipids that are eventually removed. II. Principle of the stain: Luxol fast blue, like alcian blue, is of the sulfonated copper phthalocyanine type, but is alcohol soluble, whereas alcian blue is water-soluble. Staining is due to lipoproteins, and the mechanism is one of an acid-base reaction with salt formation; the base of the lipoprotein replaces the base of the dye. Holmes attributed inconsistent results obtained with the Bodian technique to the fact that the Protargol solution never reaches the alkalinity necessary for optimal impregnation, and he modified the technique by developing a buffered impregnating solution. The pyridine in the solution is an alkali, and Holmes thought that this modified the electrostatic condition of the tissue. This is an argyrophil silver method, requiring that chemical reduction be used. The purposes of gold chloride, oxalic acid, and sodium thiosulfate are identified in the description of the Bodian procedure. III. Fixatives: 10% Neutral buffered formalin. IV. Sectioning: Cut paraffin section between 10 to 15 microns. V. Controls: A section of cerebral cortex. A spinal cord is NOT good for this procedure due to the cross section of the axons. VI. Reagents: 1. 0.1% Luxol fast blue: Luxol fast blue MBSN, 95% alcohol: Dissolve the dye in alcohol, then add: 10% Acetic acid. 2. 0.05% Lithium carbonate: Lithium carbonate& Distilled water. 3. 70% Alcohol solution: Absolute alcohol & Distilled water. 4. 10% Acetic acid: Acetic acid & Distilled water. 5. Aqueous Silver Nitrate, 20%: Silver nitrate & Distilled water 6. Aqueous Silver Nitrate, 1 %: Silver nitrate: 20% Silver solution & Distilled water 7. Boric Acid Solution: Boric acid & Distilled water 8. Borax Solution: Sodium borate & distilled water 9. 10% Pyridine Solution: Pyridine & Distilled water 10. Impregnating Solution a. Boric acid solution (fresh) b. Borax solution (fresh) c. Distilled water d. Silver nitrate, 1 % aqueous solution e. Pyridine, 10% aqueous solution Mix boric acid solution and borax solution in a 500-mL flask. Add the water, then the aqueous silver nitrate, and then the aqueous solution of pyridine. Mix thoroughly. Make enough solution for 20 mL per slide.

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11. Reducing Solution: Make fresh before use a. Hydroquinone b. Sodium sulfite (crystals) c. Distilled water 12. Gold Chloride, 0.2%: Gold chloride, 1 % solution & Distilled water 13. Oxalic Acid Solution, 2%: Oxalic acid & Distilled water 14. Sodium Thiosulfate (Hypo): Sodium thiosulfate & Distilled water. VII. 1. 2. 3. 4. 5.

Procedure: Deparaffinize sections and hydrate to distilled water. Place sections in 20% silver nitrate in the dark at room temperature for 1 hour. Prepare the impregnating solution. Take slides from 20% silver nitrate and wash for 10minutes in three changes of distilled water. Place slides in impregnating solution, allowing at least 20 mL of solution per slide. Cover jar and incubate overnight at 37°C. 6. Remove slides, shake off superfluous fluid, and place in the reducer for not less than 2 minutes. 7. Wash in running water for 3 minutes, the rinse slides in distilled water. 8. Tone in 0.2% aqueous gold chloride for 3 minutes. This solution may be reused until a brown precipitate forms or the solution becomes cloudy. 9. Rinse in distilled water. 10. Place slides in 2% aqueous oxalic acid for 3 to 10 minutes. When the axons are thoroughly blue-black, stop the process. 11. Rinse in distilled water. 12. Place the slides in 5% aqueous sodium thiosulfate. 13. Wash in tap water for 10 minutes. 14. Place the slides briefly in 95% alcohol briefly. 15. Place slides in Luxol fast blue solution and leave overnight at 56°C to 58°C. The container should be tightly capped as this alcoholic solution will evaporate readily. 16. Rinse sections in 95% alcohol to remove excess stain. 17. Place slides in distilled water. 18. Place the slides in 0.05% lithium carbonate for 15 seconds. 19. Differentiate slides in 70% alcohol for 20 to 30 seconds. 20. Rinse in distilled water. Repeat steps 18 to 20 if Luxol fast blue needs more differentiation. 21. Dehydrate in two changes each of 95% and absolute alcohols. 22. Clear in xylene and mount with synthetic resin.

VIII. Results: Myelin sheath. . . . . . . . . . . . ………………………………………………………..Blue to green Axon and nerve fibers……………………………………………………..………...Black

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Stain

Bodian Method

Holmes Silver Nitrate PTAH Method

Holzer’s Method

Fixative

10% Neutral Buffered Formalin

10% Neutral Buffered 10% Neutral Buffered Formalin Formalin

10% Neutral Buffered Formalin

Section

6-8µ

10-15µ

6-8µ

6-8µ

Control

Peripheral nerve or cerebral cortex Protargol™ Hydroquinone Formaldehyde Gold chloride Hypo Hydrochloric acid Nitric acid Aniline blue Oxalic acid Phosphomolybdic acid

Cerebral cortex

Cerebral cortex

Cerebral cortex

Silver Nitrate Boric acid Sodium borate Pyridine Hydroquinone Sodium sulfite Gold chloride Oxalic acid Sodium hyposulfite

Phosphomolybdic acid Hematoxylin 95% Alcohol Phosphotungstic acid Chloroform Lugol’s Iodine Crystal violet Oxalic acid Potassium bromide Sodium hyposulfite Potassium permanganate Aniline oil Ammonium hydroxide

Nerve Fibers – Black Background – Blue Nuclei - Black

Glial fibers – Blue Axons – Black Nerve Fibers – Black Nuclei-Blue Neurofibrils - Black Neuron-salmon Myelin-Blue

Reagents

Results

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Glial fibers – Blue

Stain

Cajal’s Stain

Weil’s Method

Luxol Fast Blue

Luxol Fast Blue/Cresyl Echt Violet

Fixative

Formalin Ammonium Bromide

10% Neutral Buffered Formalin

10% Neutral Buffered Formalin

10% Neutral Buffered Formalin

Section

10-30µ frozen sections

10-15µ

10-15µ

10-15µ

Control

Cerebral cortex

Spinal cord or Medulla

Spinal cord or Medulla

Spinal cord or Medulla

Reagents

Ammonium bromide Formaldehyde Gold chloride Mercuric chloride Hypo

Ferric ammonium sulfate Luxol fast blue Acetic acid Hematoxylin Lithium carbonate Absolute alcohol Absolute alcohol Sodium borate Potassium ferrocyanide

Luxol fast blue Acetic acid Lithium carbonate Absolute alcohol Cresyl echt violet

Results

Astrocytes- Black

Myelin Sheaths-blue to Myelin-Blue blue-black

Myelin – Blue Nissl Substances-violet Nuclei-violet

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