Selection for Electron Microscopy of Specific Areas in Large Epoxy Tissue Sections

Tissue blocks 2 × 2 × 0.4 cm were fixed 6-24 hr in phosphate-buffered 6% glutaraldehyde then sliced to 2 × 2 × 0.1 cm and rinsed in phosphate buffer for at least 12 hr. Fixation was continued for 2 hr in phosphate-buffered 1-2% OsO₄. The slices were dehydrated, infiltrated with Araldite, and embedded in flat-bottomed plastic molds. Sectioning at 4-8 μ with a sliding microtome was facilitated by addition of 10% dibutylphalate to the standard epoxy mixture. The sections were spread on water and attached to coverslips by drying, then heating to 80 C for 1 min. Staining 2 min with 1-3% KMnO₄ and temporary mounting in glycerol on a slide allowed the desired area for electron microscopy to be selected and marked. This area was then cemented to the facet of a conventional epoxy casting with a drop of epoxy resin (without added dibutylphthalate). After polymerization, the coverslip was removed by quick cooling leaving a flat re-embedded portion of the original section. This portion was viewed by transillumination in a dissecting microscope and trimmed of surplus tissue. Ultrathin sections for electron microscopy were obtained in the usual manner.     

Acceleration of Mallory's Phosphotungstic Acid-Hematoxylin Staining for Skeletal Muscle Fixed in Formalin

The staining time for mammalian skeletal muscle fixed in neutral phosphate-buffered formalin was shortened from 12-24 hr to 10-30 min. The permanganate-oxalate sequence was omitted although oxidation by periodic acid or with iodine was found to be necessary. The material was embedded in paraffin and cut 6 μ or less. Deparaffinized sections were treated with 1% alcoholic iodine for 10 rain followed by 5% Na₂S₂O₃ for 2 min and placed in an oven at 60 C for 10-30 min to stain in a preheated mixture of 50 ml of ripened Mallory's phosphotungstic acid-hematoxylin and 1 ml of 2% phosphomolybdic acid. Experiments with fixation showed that the staining procedure followed Zenker's fluid successfully but not Bouin's fluid. Oxidation by KMnO₄ was effective only after Zenker fixation; oxidation by CrO₃ was unsuccessful.     

Staining Pericellular Boutons of Glutaraldehyde-Fixed Nervous Tissue; A Chromation-Silvering Sequence for Frozen Sections

After fixing in phosphate-buffered 5% glutaraldehyde, pH 6.8, by perfusion, brains were sliced to 3-5 mm pieces which were placed in the fixative for 5-7 days. The pieces were washed through several changes of 2.26% NaH₂PO₄ for 12 hr, 30 μ frozen sections cut, and mordanted 2 days in an equal-parts mixture of 3.5% CrO₃ and 5% Na-tartrate, which had been aged at 20-25 C for 20 days prior to use. After washing in distilled water, the sections were put into a solution containing AgNO₃, 20 gm; and KNO₃, 15 gm, in distilled water, 80 ml; at 30 C for 1.5-2 hr, then reduced at 40-45 C in three pyrogallol solutions as follows: 1-2 sec in 1% pyrogallol in 55% alcohol; 3-4 sec in a 0.67% solution in 33% alcohol, and 5-7 sec in a 0.5% solution in 25% alcohol. Gold toning is optional; dehydration, clearing and covering, routine. The technic shows particularly the perisomatic fibers, boutons en passant and boutons termineaux. Fibers in nerve tracts may be visible but lightly stained; cell nuclei may be dark, but the cytoplasm remains pale.     

A Phloxine-Azure-Hematoxylin Sequence for Differential Staining of Cells in Pancreatic Islets

Sections of 6 μ from tissues fixed in Susa or in Bouin's fluid (without acetic acid) and embedded in paraffin were attached to slides with Mayer's albumen, dried at 37 C for 12 hr, deparaffinized and hydrated. The sections fixed in Susa were transferred to a I₂-K₁ solution (1:2:300 ml of water); rinsed in water, decolorized in 5% Na₂S₂O₃; washed in running water, and rinsed in distilled water. Those fixed in Bouin's were transferred to 80% alcohol until decolorized, then rinsed in distilled water. All sections were stained in 1% aqueous phloxine, 10 min; rinsed in distilled water and transferred to 3% aqueous phosphotungstic acid, 1 min; rinsed in distilled water; stained 0.5 min in 0.05 azure II (Merck), washed in water; and finally, nuclear staining in Weigert's hematoxylin for 1 min was followed by a rinse in distilled water, rapid dehydration through alcohols, clearing in xylene and covering in balsam or a synthetic resin. In the completed stain, islet cells appear as follows: A cells, purple; B cells, weakly violet-blue; D cells, light blue with evident granules; exocrine cells, grayish blue with red granules.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

Triple Silver Impregnation for Selective Staining of Avian Nerves

Frozen sections of avian tissue fixed 7 days or longer in 10% formalin or formol-saline are cut at 20-50 μ, left in distilled water for 2 hr, and placed in 0.002% aqueous AgNO₃ for 3-4 days. Subsequent procedure is essentially that of Weddell and Glees. Sections are placed in 20% AgNO₃ for 30 min, then carried through 3 baths of 3% formalin in less than 10 min. Immediately thereafter they are washed 1-2 sec in a 0.1% solution of NH₄OH (cone) and placed in the ammoniacal silver solution (made with 20% AgNO₃) until the nerves become distinct, as seen under a microscope; usually, in about 15 min. After washing briefly, the sections are fixed in 5% Na₂S₂O₃ for 3-10 min, dehydrated, cleared, and mounted in the usual way.     

Staining Procedures for Ultrathin Sections of Tissues Embedded in Polyester Resin

Tissues were fixed for 30 min In cold (0-2° C) 1% OsO₄ (Palade) buffered at pH 7.7, to which 0.1% MgCl₂ was added. Dehydration was in a graded ethanol series (containing 0.5% MgCl₂) at 0-2° C, and terminated with 2 changes of absolute ethanol. Tissues were then transferred by a graded series to anhydrous acetone. Infiltration of the tissue with Vestopal-W (a polyester resin), is gradual with the aid of graded solutions of Vestopal-W in acetone. The infiltrated tissue is encapsulated and initial polymerization is done under ultraviolet light at room temperature for 8-16 hr. This is followed by final hardening at 60° C for 36-48 hr. Sections (0.2-1 μ) were cut, dried on slides, placed in acetone for 1 min and then treated by either of the following staining procedures: (1) Thionin-azure-fuchsin staining: Flood the preparation with 0.2% aqueous thionin and heat to 60-80° C for 3 min; if the preparation begins to dry, add stain. Rinse in distilled water. Flood the slide with 0.2% azure B in phosphate buffer at pH 9. Heat to 60-80° C for 3 min; do not permit the preparation to dry. Rinse in distilled water. Dip the slide in MacCallum's variant of Goodpasture's carbol-fuchsin stain for 1-2 sec. Rinse in distilled water. Check the preparation microscopically for intensity of the fuchsin stain. Repeat dips as may be needed to obtain the desired intensity. Rinse in distilled water. Dehydrate quickly in 95% and absolute alcohol; clear in 2 changes of xylene and cover in Permount or similar synthetic resin. (2) Thionin-azure counterstain for the periodic acid-Schiff reaction: Oxidize the tissue in 0.5% periodic acid for 15 min and transfer to Schiff's leucofuchsin solution for 30 min. Counterstain with 0.5% aqueous thionin for 3 min; wash in distilled water; stain in 0.2% azure B in phosphate buffer at pH 5.5; wash in distilled water; dehydrate; clear and cover as in the first method. For temporary preparations let dry after absolute alcohol and apply a drop of immersion oil directly on the section.     

Differentiation of Cells in the Hypophysis and in Pancreatic Islets

Deparaffinized, 3-5μ, sections are brought to water, oxidized 3.5 min in an equal-parts mixture of 0.3% H₂SO₄ and 0.3% KMnO₄, and decolorized with 4% K₂S₂O₅. Nuclei are stained with Gomori's (1939) chromium-hematoxylin, and cell granules with Cason's (1950) mixture. The eosinophilic cells of the hypophysis and the alpha cells of pancreatic islets (of Langerhans) stain carmine red; basophilic and beta cells stain dark blue. Heidenhain's _susa is the most suitable fixative for hypophysis, Bouin's fluid for pancreas; but a satisfactory result is obtainable after formalin-sublimate or plain formalin. Besides studying the ratio of the cell types in the hypophysis or in pancreatic islets, it is possible to estimate the granule content of the cells. The method works on human autopsy material provided fixation of hypophysis occurs within 24 hr, and. pancreas, 12 hr post mortem, and it is suitable also for quite fresh organs.     

Enhanced Reliability in Silver Impregnation of Terminal Axonal Degeneration-Original Nauta Method

Brains of rat with surgical lesions 3-5 days old are fixed in 10% neutralized formalin (excess of CaCO₃), 20 μ serial frozen sections cut therefrom and kept in neutralized formalin for an additional 24-48 hr. The sections are soaked in distilled water 12-24 hr, transferred to 50% alcohol containing 0.75 ml of concentrated NH₄OH (sp. gr. 0.91) per 100 ml 12-24 hr, placed in distilled water 2-3 hr and then in silver-pyridine solution (AgNO₃ 3% aq., 20 ml; pyridine, 1 ml) for 48 hr. Test sections are transferred directly to each one of 3 ammoniated silver-solutions, pH 12.8, 13.0 and 13.2, made as follows: To 200 ml of solution 1 (silver nitrate, 6.4 gm; alcohol 96%, 220 ml; NH₄OH (sp. gr. 0.91), 28 ml and distilled water, 440 ml) is added respectively 8-12 ml, 12-16 ml and 16-20 ml of solution 2 (2% NaOH) to give the pH desired. The test sections are studied and the optimal ammoniated silver solution chosen. Two baths of ammoniated silver are used, the section placed with continuous agitation into the first bath for 30 sec and the second bath for 60 sec. The sections are then transferred directly into a reducing bath (formalin 10%, 2ml; alcohol 96%, 5 ml; citric acid 1%, 1.5 ml and distilled water, 4.5 ml) for 2 min and from there to 5% Na₂S₂O₃ for 1 min, rinsed in 3 changes of distilled water, dehydrated and mounted.     

A HYDROXYL-DEPENDENT SILVER METHOD FOR NERVE AXOPLASM

Axoplasm is selectively impregnated by the following steps: (1) fixation in 10% formalin or in 10% formalin with added sucrose, 15%, and concentrated NH₄OH, 1%, for 1-7 days; (2) frozen sections; (3) extraction of the sections in 95% ethyl alcohol, absolute alcohol, xylene, and 95% ethyl alcohol and absolute alcohol, 1 hr each; (4) distilled water, 3 changes of 10 min each; (5) 20% AgNO₃ (aq.) at 25°C, 30 min; (6) distilled water, 3 changes of 1-2 sec each; (7) 6.9% K₂CO₃, 1 hr; (8) water, 3 changes of about 1 min each; (9) 0.2%AuCl₃, 2 min; (10) distilled water; (11) 5% Na₂S₂O₃, 2 min; (12) washing, clearing and mounting. This procedure is proposed as a simplified stain for axoplasm, with other tissue components remaining unstained. The few reagents necessary suit this method for histochemical investigation of the mechanism of silver staining.     

Epon-Embedded Tissue Stained with Aldehyde-Fuchsin for Radioautography

Four-week-old Holtzman rats were injected intraperitoneally with 20 μCi ¹²⁵I. Six or eight weeks later, they were killed by intracardiac perfusion with glutaraldehyde; thyroid and adrenal glands were excised, postfixed in osmic acid, and embedded in Epon. Steps in the staining procedure of 0.5-1 μm thick sections are: oxidation in 0.3% potassium permanganate in 0.625% sulfuric acid, 2-5 min at 70 C; brief rinse; bleaching with 2.5% NaHSO₃, 4-5 min; brief r-utse; let dry completely; aldehydefuchsm, 15-20 min at 50 C; 95% alcohol; rinse in absolute alcohol; let dry completely. seaions were coated with Kodak NTB2 emulsion and exposed for 3 to 8 weeks. Results indicate that (1) tissues are well stained even after an 8-week exposure, (2) aldehyde-fuchsin pduces no chemographic effect, and (3) structures underneath the emulsion are easily identified.     

Inhibition of antennal esterases of the egyptian armyworm Spodoptera littoralis by trifluoromethyl ketones

A series of aliphatic and aromatic trifluoromethyl ketones has been tested as inhibitors of the antennal esterases of the Egyptian armyworm Spodoptera littoralis, by evaluation of the extent of hydrolysis of [1-³H]-(Z,E)-9, 11-tetradecadienyl acetate (1), a tritiated analog of the major component of the sex pheromone. The most active compounds with a long chain aliphatic structure were 3-octylthio-1,1,1-trifluoropropan-2-one (2) (IC₅₀ 0.55 μM) and 1,1,1-trifluorotetradecan-2-one (4) (IC₅₀ 1.16 μM). The aromatic compounds were generally less potent inhbitors than the coressponding aromatic ones, although β-naphthyltrifuloromethyl ketone (10) exhibited a remarkable inhibitory activity (IC₅₀ 7.9 μM). Compounds 2, 4 and 10 exhibit a competitive inhibition with K_i values of 2.51×10⁻⁵ M, 2.98×10⁻⁵ M and 2.49×10⁻⁴ M, respectively. Some of the trifluoromethyl ketones tested were slow-binding inhibitors and compounds 2 and 10 are described as inhibitors of the antennal esterases of a moth for the first time.     

Azure a-Schiff, Alcian Blue, HIO4-Schiff, Naphthol Yellow S—A Sequential Staining Method for Paraffin Sections

Paraffin sections from tissue fixed 4-12 hr in 10% formalin containing 0.5% cetyl pyridinium chloride, and washed 2 hr, were stained as follows: (1) Hydrolyze in 5 N HCl at room temperature for 8.5-9 min, or use standard Feulgen hydrolysis at 60 C. (2) Stain in azure A-Schiff, 0.5% in bisulfite bleach (1 N HCl, 5; 10% Na₂S₂O₅, 5; and distilled water 90—parts by volume) for 10 min. (3) Place in bisulfite bleach 2 changes, 2 min each; wash in water, 1-2 min. (4) Stain in Alcian blue (0.1% in 0.01 2V HCl, pH 2.0) for 10 min. (5) Place in 0.01 N HCl for 2-3 min; wash in water for 1-2 min. (6) Oxidize in 0.5% HIO₄ for 5 min; wash in water, 1-2 min. (7) Stain in Schiff's leucofuchsiu, 10 min. (8) Treat with bisulfite bleach as in step 3; wash in running water, 10 min. (9) Stain in naphthol yellow S (0.01% in 1% acetic acid) for 1-2 min. (10) Place in 1% acetic acid for 2 min, dehydrate in tertiary butanol, clear and cover. Result: DNA is deep blue; acidic mucins are light blue; neutral polysaccharides, red to magenta; and proteins, yellow. Proper timing of the hydrolysis for the Feulgen reaction is the most critical step. Overhydrolysis results in green nuclei (staining by naphthol yellow S) whereas purplish nuclei are the results of insufficient hydrolysis.     

Epoxy Embedding, Sectioning and Staining of Plant Material for Light Microscopy

By using a formula which gives a relatively soft epoxy embedding medium, it is possible to cut sections of plant material with a sliding microtome equipped with a regular steel knife. Blocks having a cutting face of 10 × 10 mm, giving sections of 4-10 μm, can be used. Tissues are fixed in Karnovsky's fluid, postfixed in 1 or 2% OsO₄, embedded in Spurr's soft epoxy resin, Araldite, or Epon mixtures. 5% KMnO₄, followed by 5% oxalic acid, then neutralized in 1% LiCO₃, are used to mordant the sections. Some of the stains used are Mallory's phosphotungstic acid-hemotoxylin, acid fuchsin and toluidine blue, or toluidine blue. Mounting is done with whichever soft epoxy resin was used in casting the blocks.     

A Copper Sulfate-Silver Nitrate Method for Nerve Fibers of Planarians

For staining in toto, planarians are fixed in a mixture of 10 ml of commercial formalin, 45 ml of 95% ethanol and 2 ml of glacial acetic acid. After treatment with 70% ethanol 3-10 days, they are washed in distilled water and immersed in 10% CuSO₄. 5H₂O for 3 hr at 50° C, transferred without washing to 1% AgNO₃ for 1.0-1.5 hr at 50° C; and then developed in: 10 ml of 1% pyrogallol, 100 ml of 56% ethanol and 1 ml of 0.2% nitric acid. Gold toning, 5% Na₂S₂O₃ and dehydration follow as usual. For staining sections, material is fixed in the same fixative, embedded in paraffin and sectioned at 10 μ. After bringing sections to water, they are immersed in 20% CuSO₄. 5H₂O for 48 hr at 37° C; then rinsed briefly in distilled water and placed in 7% AgNO₃ for 24 hr at 37° C. They are washed briefly in distilled water and reduced in: hydroquincne, 1 gm; Na₂SO₃, 5 gm and distilled water 100 ml. Gold toning, followed by 5% Na₂S₂O₃ and dehydration completes the process. Any counterstaining may follow.     

A Glutaraldehyde-Dichromate-Silver Sequence for Differentiating Norepinephrine Cells from Epinephrine Cells in Neonatal and Adult Rats—Paraffin Sections

A glutaraldehyde-K₂Cr₂O₇ procedure intensified by silver staining enabled norepinephrine and epinephrine cells to be distinguished readily in paraffin sections of the adrenal glands of rats 8 days after birth. The technique involved fixation in 0.1 M cacodylate-buffered 5% glutaraldehyde (6-24 hr), treatment with 3.5% K₂Cr₂O₇ (6-12 hr) and routine preparation of paraffin sections. The sections were deparaffinised, brought to water and immersed in Fontana's solution (24 hr), prepared by adding concentrated NH₄OH drop by drop to 5% AgNO₃ until the precipitate formed just redissolved; more 5% AgNO₃ was then added until a permanent cloudiness just developed. After a rinse in distilled water, the sections were treated with 0.5% gold chloride (5 min) and Na₂S₂O₃ (5 min), then mounted in Depex. This sequence resulted in an intense black cytoplasmic colouration in norpinephrine-containing cells of both the adult and 8-day-old animals whereas epinephrine-containing cells remained colourless. The glutaraldehyde-K₂CrO₇ procedure, without intensification, gave very clear results in the adult: a yellow cytoplasmic colour in the norepinephrine cells with epinephrine cells colourless. A glutaraldehyde-OsO₄ sequence gave a less well defined separation of these cell types in the adult and failed to distinguish the cell types in the neonate.     

Staining Residual Lipids in Ultrathin Sections of Tissues Embedded in Polyester Resin

Rat suprarenal glands fixed in Palade's 1% OsO₄, buffered at pH 7.7 with veronal-acetate, to which 0.1% MgCl₂ was added, were embedded in Vestopal-W and sectioned at 0.2-1 µ. The sections were attached to slides by floating on water, without adhesive, and drying at 60-80° C, placed in acetone for 1 min and then treated with the following staining procedure: Place the preparation in a filtered solution of oil red O, 1 gm; 70% alcohol, 50 ml; and acetone, C.P., 50 ml; for 0.5-1 hr. Rinse in absolute ethyl alcohol; drain; counterstain with 0.5% aqueous thionin for 5 min; rinse in distilled water; drain; stain in 0.2% azure B in phosphate buffer at pH 9, for 5 min. Dry and apply a drop of immersion oil directly on the section. The preparations are temporary. Ciaccio-positive lipids, rendered insoluble by OsO, fixation, stained red to ochre.     

Correlation of CrystaL Growth with the Staining of Axons by the Golgi Procedure

The surface of the specimens subjected to a modified Golgi technique (formalin fixed material; specimens in the following solution for 8-10 days at 27 C: 3% K₂Cr₂O₇, 100 ml, with the addition of 2.5-10 ml of 10% formalin and 6-25 gm of sucrose; then in 0.75% AgNO₃ for at least 2 days at 27 C) is sometimes covered with a fur of filamentous crystals and sometimes with a powdery precipitate of laminar crystals. In a series of experiments in which about 500 blocks of tissue were treated with variations of the staining procedure, good axonal stain was positively correlated with the appearance of filamentous crystals. These filaments have a thickness of 1-4 μ and grow at a rate of 160-330 μ/hr, reaching a length of 2-7 mm.     

Fluorescence in Paraffin Sections of Dye-Labelled Protein Administered in vivo: Effect of HgCl2 Fixation

Lung and liver slices, 2-3 mm thick, from guinea pigs injected intravenously with fluorescent dye-protein conjugate are fixed for 15-30 min in saturated aqueous HgCl₂, dehydrated in ethanol, cleared in xylene and embedded in paraffin at 60 C. Mercurial deposits are removed with I₂KI from 5 μ sections taken to water, and the iodine then removed with 5% Na₂S₂O₃. Sections are mounted from xylene into permanent nonfluorescent mounting medium. This procedure gives optimal fluorescence which is not decreased by the technic of removing mercurial precipitates. Longer fixation, fixation in phosphate-buffered formalin, or in an HgCl₂-formalin mixture gives inferior results.     

A Sensitive Myelin-Sheath Stain Particularly Useful for Small Mammals and Neonatal Cats

Staining of myelinated fibers including the delicate myelin sheaths of infantile animals is as follows: perfuse the anesthetized animal with a pH 7.4 posphate-buffered fixative, either 10% formalin, 6% gluteraldehyde or a mixture containing 3% gluteraldehyde and 2% acrolein. Dissect out the brain or spinal cord and continue fixation for at least 24 hr. Cut larger brains to 1 cm in at least one dimension. Wash in running tap water 2-3 hr and soak in 2.5% potassium dichromate in 1% acetic acid (the primary mordant) for 3-5 days in darkness. Wash at least 12 hr in running tap water. Dehydrate and embed in celloidin and store in 80% ethanol. Section at 25-60 μ into 80% ethanol. Wash 1-2 min in distilled water and then immerse in 1-2% ferric alum at 50 C for at least 1 hr (the secondary mordant). Wash in tap water and stain at least 1 hr at 50-60 C in 0.5% unripened hematoxylin in 1% acetic acid. Wash well in tap water and differentiate in a mixture containing 0.5% ferrityanide, 0.5% borax and 0.5% Na₂CO₃; 2 changes. Wash well in distilled water, then in tap water, and dehydrate, clear and mount. Myelin stains black, cell bodies stain tan, and the background is pale yellow. With minor modifications in timing, the method is applicable to frozen and to paraffin sections; the primary mordant being omitted in the freezing technique.