The Demonstration of Beta Cells in Pancreatic Islets and Their Tumors

The stain is applied routinely to tissues fixed in 10% buffered formalin (pH near 7.0) or in Bouin's fluid. Bring paraffin section to water as usual and mordant 72 hr in 5% CrCl₃ dissolved in 5% acetic acid. Wash in water and in 70% alcohol and stain 6 hr. Formula of staining solution: new fuchsin, 1% in 70% alcohol, 100 ml; HCl, conc., 2 ml and paraldehyde, 2 ml, mixed together and added to the dye solution; let stand 24 hr before use. After staining, wash in running tap water 5-10 min, rinse in distilled water and counterstain if desired. Dehydration in alcohol, clearing and covering completes the process. When the paraldehyde is obtained from a freshly opened bottle, standardized staining times can be used and thus eliminate the necessity of differentiating individual slides. The granules of beta cells stained deep blue to purple and were demonstrated in the pancreatic islet of man, dog, mouse, frog, guinea pig and rabbit.     

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).     

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.     

Use of Gallocyanin as a Myelin Stain for Brain and Spinal Cord

Tissue fixed in 10% formalin, formol saline, CaCO₃ or phosphate buffer neutralized formalin, Baker's formol calcium, Cajal's formol ammonium bromide, formalin-95% ethanol 1:9, formalin-methanol 1:9, Lillie's methanol-chloroform or Salthouse's formol cetyltrimethylammonium bromide was dehydrated and embedded in paraffin. Sections were attached to slides with either albumen or gelatine adhesive and processed throughout at room temperature of 22-25 C. Mordanting 30-60 min in 1% iron alum was followed by a 10 min wash in 4 changes of distilled water. Myelin was stained in a gallocyanin self-differentiating solution for 1-2.5 hr; thick sections requiring the longer time. The staining solution (pH approximately 7.4) consisted of Na₂CO₃, 90 mg; distilled water, 100 ml; gallocyanin, 250 mg; and ethanol, 5 ml. The ethanol was added to this mixture last, and after the other ingredients had been boiled and then cooled to room temperature. After a staining and thorough washing, Nissl granules were stained for 5-10 min in a solution consisting of: 0.1 M acetic acid, 60 ml; 0.1 M sodium acetate, 40 ml; methyl green, 500 mg. Washing, dehydration, clearing and mounting completed the process. Myelin sheaths were stained dark violet; neuronal nuclei, light green with dark granules of chromatin; nucleoli of motor cells and erythrocytes, dark violet; cytoplasm, green with dark green Nissl granules. The simple and reliable method can be adapted easily for use with automatic tissue processors.     

Silver Impregnation of Degenerating Axons in the Central Nervous System: A Modified Technic

Frozen sections of formalin-fixed brains containing surgical lesions, were treated with 15% ethanol for 0.5 hr., soaked in 0.5% phosphomolybdic acid for 0.25-1.0 hr., and subsequently treated with 0.05% potassium permanganate for 4-10 min. (The duration of the latter treatment is critical and individually variable). Subsequent procedure is as follows: decolorize in a mixture of equal parts of 1% hydroquinone and 1% oxalic acid; wash thoroughly and soak sections in 1.5% silver nitrate for 20-30 min.; ammoniacal silver nitrate (silver nitrate 0.9 g., distilled water 20 ml., pure ethanol 10 ml., strong ammonia 1.8 ml., 2.5% sodium hydroxide 1.5 ml.) 0.5-1.0 min.; reduce in acidified formalin (distilled water 400 ml., pure ethanol 45 ml., 1% citric acid 13.5 ml., 10% formalin 13.5 ml.) 1 min.; wash, and pass section through 1 % sodium thiosulf ate (0.5-1.0 min.); wash thoroughly and pass sections through graded alcohols and xylene (3 changes); cover in neutral synthetic resin.     

A Six-Dye Staining Schedule for Sections of Mesquite and Other Desert Plants

Materials are fixed in FPA (formalin, 2; propionic acid, 1; 70% ethanol, 17). Paraffin sections on slides are brought to 50% ethanol and stained as follows: (1) in Bismarck brown Y, a 0.02% solution in 0.1% aqueous phenol, 10-30 min; wash 30 sec in 0.7% acetic acid, and wash in distilled water 20-30 sec; (2) in crystal violet, 1% in 70% ethanol alkalinized with 1 drop of 1 N NaOH per 100 ml, 12-35 min; wash 30-60 sec in tap water to remove excess stain, and rinse 0.5 sec in 70% ethanol; then mordant in I₂-KI, 1% each in 70% ethanol, 40 sec, and rinse in 70% ethanol 2-5 sec; (3) in a mixture containing 0.4% acid fuchsin and 0.6% crythrosin B in 70% ethanol about 0.5 sec; rinse in 70% ethanol 5-15 sec to remove excess red; dehydrate in 70%, 95%, and absolute ethanol, 2-3 sec each; (4) in fast green FCF, 0.5% in a mixture of equal parts of methyl cellosolve, absolute ethanol, and clove oil, 5-15 sec; rinse in a mixture of clove oil, 10 ml; absolute ethanol, 100 ml; and methyl cellosolve, 10 ml, 5-7 sec; (5) in orange G, 0.75 gm in a mixture of clove oil, 40 ml; absolute ethanol, 40 ml; and methyl cellosolve, 60 ml, 5-30 sec; rinse clean in a 1:1 mixture of xylene and absolute ethanol, 5-20 sec Complete the clearing in pure xylene, 3 changes, 1.5 min in each, and apply a cover glass with synthetic resin. Slides are agitated in all steps except Bismark brown Y, crystal violet, and the xylenes. Contrast and staining intensity are adjusted by varying staining times in the dye solutions.     

Staining of Small Lymphoid Nucleoli in Paraffin Sections by Aniline-Azure B

To see small lymphoid nucleoli clearly in 1-2 μ paraffin sections, the staining of contiguous chromatin masses in the nucleus was suppressed by a hydrolysis-aniline blocking sequence, which produces aldehyde from DNA, and attaches aniline to that aldehyde to make a diphenamine base, thus reducing the acidity of the chromatin and its affinity for basic dyes. Nucleolar RNA remains fully stainable by azure B, because the hydrolysis used does not produce aldehyde groups in it, to allow aniline attachment. Technique: Hydrolyse the 10% formol-saline fixed, deparaffinised 1-2 μ section for 4.5-5.0 min in 10% (v/v) HCl in tetra-hydrofuran at 39-40 C, rinse in water, and treat at room temperature in 10% (v/v) aniline in acetic acid for 10 min. Stain 2-4 hr with freshly prepared 0.1% azure B in a 1:10 dilution of tris buffer at pH 7.0. Rinse, blot off excess water, pass through acetone and xylene to a polystyrene mounting. DNA stains pale green to colourless; nucleolar and cytoplasmic RNA, blue.     

Staining Nerve Fibers After Sublimate-Acetic and After Bouin'S Fluid

A silver staining method for paraffin sections of material fixed in HgCl₂, sat. aq., with 5% acetic acid is as follows. Process the sections through the usual sequence of reagents, and including I-KI in 70% alcohol, thiosulfate (5% aq.), washing and back to 70% alcohol containing 5% of NH₄OH (conc. aq.). After 3 minutes in the ammoniated alcohol, wash through tap water and 2 changes of distilled water and silver 5-10 minutes at 25°C. in 15% AgNO₃ aq. to which 0.02 ml. of pyridine per 100 ml. has been added. Blot the slide, but not the section and do not rinse. Reduce at 45°C. in 0.1% pyrogallol in 55% alcohol, then rinse in 55% alcohol and wash in water. The remainder of the process consists of gold toning, intensifying in oxalic acid, fixing in 5% Na₂S₂O₃, washing, dehydrating, clearing and covering. When the specimen contains much smooth muscle, the I-KI solution is acidified before use by adding 2 ml. of 1N nitric acid per 100 ml., and the sections treated for 3 minutes instead of the usual 2 minutes. Formalin should not be added to sublimate-acetic, but specimens that do not contain strongly argyrophilic nonneural tissue may be fixed in formalin or, preferably, Bouin's fluid. Sections of tissue after the latter type of fixation will not require the I-KI and thiosulfate but can go from 95% alcohol to the ammoniated alcohol. The advantages of fixing in HgCl₂-acetic acid are suppression of the staining of connective tissue and intensifying the staining of nerve fibers.     

A modification of the tannic acid-phosphomolybdic acid-dye stain for demonstrating myoepithelial cells in formalin fixed tissue

A modified tannic acid-phosphomolybdic acid-dye procedure is used for staining myoepithelial cells in formalin fixed surgical and autopsy material. Paraffin sections are brought to water, mordanted for 1 hr in Bouin's fixative previously heated to 56 C, cooled while still in Bouin's, rinsed in tap water until sections are colorless, rinsed in distilled water, treated with 5% aqueous tannic acid 5-20 min, rinsed in distilled water 30 sec or less, treated with 1% aqueous phosphomolybdic acid 10-15 min, rinsed 30 sec in distilled water, rinsed in methanol, stained 1 hr in a saturated solution of amido black or phloxine B in 9:1 methanol:acetic acid, rinsed in 9:1 methanol:acetic acid, dehydrated, cleared and mounted. Myoepithelial cells of sweat, lacrimal, salivary, bronchial, and mammary glands are blue-green with amido black or pink with phloxine B. Fine processes of myoepithelial cells are well delineated. Background staining is minimal and the procedure is highly reproducible.     

A Modification of the Tannic Acid Phosphomolybdic Acid-Dye Stain for Demonstrating Myoepithelial Cells in Formalin Fixed Tissue

A modified tannic acid-phosphomolybdic acid-dye procedure is used for staining myoepithelial cells in formalin fixed surgical and autopsy material. Paraffin section are brought to water, mordanted for 1 hr in Bouin's fixative previously heated to 56 C, cooled while still in Bouin's, rinsed in tap water until sections are colorless, rinsed in distilled water, treated with 5% aqueous tannic acid 5-20 min, rinsed in distilled water 30 sec or less, treated with 1% aqueous phosphomolybdic acid 10-15 min, rinsed 30 sec in distilled water, rinsed in methanol, stained 1 hr in a saturated solution of amido black or phloxine B in 9:l methanol:acetic acid, rinsed in 9:l methanol:acetic acid, dehydrated, cleared and mounted. Myoepithelial cells of sweat, lacrimal, salivary, bronchial, and mammary glands are blue-green with amido black or pink with phloxine B. Fine processes of myoepithelial cells are well delineated. Background staining is minimal and the procedure is highly reproducible.     

Tannic acid, Iron hematoxylin, Alcian blue, and Basic fuchsin for staining islets and reticular fibers of the pancreas

In this technique alpha cells are stained by basic fuchsin, beta cells by iron-hematoxylin, reticular fibers by ferric tannate, and much by alcian blue. Among 6 commonly used fixatives tested, Bouin's fluid fixation (8-12 hr) gave the best staining results. Procedure: paraffin sections to water; 0.5% Li₂CO₃ to remove picric acid; 20% tannic acid, 15 min; wash well; 2-4 sec in 0.5% basic fuchsin containing 10% alcohol; rinse, then differentiate in 1% aniline in 90% alcohol until alpha cells are red and beta cells pink; 1% phosphomolybdic acid, 1 min; 5% hematoxylin in 2% iron alum, 0.5 min; wash well; 1% filtered alcian blue SGX, 15 sec; rinse, dehydrate, clear, and mount in synthtic resin. Results: reticular fibers, black; acinar cells, orange to gray; alpha cells, red; collagenous fibers, red; beta cells, gray granules; ducts, bluish-green. The method was tested on rat, rabbit, dog, hamster, cow and man.     

A Combined Hcl-Acetic-Alcohol Fixation and Hydrolysis Followed by Cresyl Violet Staining for Mosquito Chromosome Spreads

Mosquito tissues of cytogenetical importance were dissected out on a slide in 0.65% NaCl, under a dissecting microscope, and treated about 30 sec in a drop of 1:3 Carnoy's fixative diluted 1:19 with distilled water. Fixing and hydrolysis was done by a single step in a mixture consisting of: glacial acetic acid, 1; ethanol 96%, 3; HCl conc., 2; and distilled water, 2 (v/v) for 2-6 min at 20-25 C. The specimen was then rinsed with the acetic-alcohol fixative and covered in a drop of 1% cresyl violet in 50% acetic acid under a coverslip coated with Mayer's albumen. Washing was performed immediately by adding water dropwise to one side of the coverslip and drawing the fluid from the other side with absorbent paper. The preparation could be used either as a temporary slide or made into a durable mount. The DNA-containing bands of the giant polytenic chromosomes stained dark violet; interband regions, weakly stained or colourless against a clear background. Mitotic and meiotic figures in gonadal cells stained selectively dark violet or violet with a practically unstained cytoplasm.     

Brazilin-Toluidine Blue O and Hematoxylin-Darrow Red Methods for Brain and Spinal Cord

Tissue fixed in 10% formalin, formalin-95% ethanol 1:s CaCO₂ or phosphate buffer neutralized formalin, or methanol-chloroform 2:1, was dehydrated and embedded in paraffin or double-embedded by infiltration in 1% celloidin followed by a chloroform-paraffin sequence. Sections were attached to slides with either albumen or gelatine adhesive and processed throughout at room temperature of 24-26 C. For either method, mordanting 30-60 min in 1% iron alum was followed by a 10 min wash in 4 changes of distilled water. For brazilin-toluidne blue O, myelin was stained for 20-60 min, depending upon section thickness, in a self-differentiating solution consisting of: 0.15% Li₂CO₃ 75 ml; 6% brazilin in 95% ethanol, 25 ml; and NaIO₃ 75 mg. After a thorough washing, Nissl material was stained for 3-8 min in a solution consisting of: 0.1 M acetic acid, 90 ml; 0.1 M sodium acetate, 10 ml; and 1% toluidine blue 0, 2.5 ml. For hematoxylin-Darrow red, myelin was stained for 2-6 hr in a self-differentiating solution consisting of: 0.15% Li₂,CO₃ 95 ml; 10% hematoxylin in 95% ethanol, 5 ml; and NaIO₃ 25 mg. After a thorough washing, Nissl material was stained for 20 min or less in a solution consisting of: 0.1 M acetic acid, 90 ml; 0.1 M sodium acetate, 10 ml; Darrow red, 25 mg. This mixture was first boiled, cooled to room temperature and filtered. In both methods, washing, dehydration, clearing, and mounting completed the process. In the brazilin-toluidine blue technic, myelin sheaths were stained reddish purple; neuronal nuclei light blue with dark granules of chromatin; nucleoli dark blue; and cytoplasm blue with dark blue Nissl granules. In the hematoxylin-Darrow red procedure, myelin sheaths were blue-black; nuclei light red with dark granules of chromatin; nucleoli almost black; and cytoplasm red with bright red Nissl granules.     

Selective Silver Impregnation of Degenerating Axons In The Central Nervous System

Rat and rabbit brains containing surgical lesions of 5-10 days' duration were fixed in 10% formalin (neutralized with calcium carbonate) for 1 week to 6 months. Frozen sections (15-20 n) were rinsed and then soaked 7 minutes in a 1.7% solution of strong ammonia in distilled water. Subsequent treatment was as follows: rinse; 0.05% aqueous potassium permanganate 5-15 minutes; 0.5% aqueous potassium metabisulfite, 2 changes of 2.5 minutes each; wash thoroughly in 3 changes distilled water; 1.5% aqueous silver nitrate, 0.5-1.0 hr.; 1% citric acid, 5-10 sec.; 2 changes distilled water; 1% sodium thiosulfate, 30 see.; 3 changes distilled water. Each section is then processed separately. Ammoniacal silver solution (450 mg. silver nitrate in 10 ml. distilled water; add 5 ml. ethanol; let cool to room temperature; add 1 ml. strong ammonia water and 0.9 ml. of 2.5% aqueous sodium hydroxide), 0.5-1.0 min. with gentle agitation. Reduction of about 1 minute is accomplished in: distilled water, 45 ml.; ethanol, 5 ml.; 10% formalin, 1.5 ml.; 1% citric acid, 1.5 ml. Rinsing; 1% sodium thiosulfate, 10 sec.; thorough washing followed by dehydration through graded alcohol and 3 changes of xylene or toluene complete the staining process. Normal nerve fibers are slightly stained to unstained, degenerating fibers, black. The treatment in potassium permanganate is critical since too little favors overstaining of normal fibers and too much abolishes staining of degenerating fibers.     

Application of the Nauta-Gygax Technic for Degenerating Axons to Mounted Sections

Frozen sections of formalin-fixed brains containing lesions were mounted on slides that had been coated first with albumen-glycerol (1:1) then 4% gelatin and blotted. The slides were placed in formaldehyde vapor at 56° C for 40-60 min, washed, and stored (optional) in 10% formalin-saline. The staining technic was as follows: after washing, soak 30-40 min in 0.5% phosphomolybdic acid, rinse; put in 0.05% potassium permanganate 9-16 min (usually 12 min); decolorize in a 1:1 mixture of 1% hydroquinone and 1% oxalic acid; wash thoroughly; soak in 1.5% AgNO₃ at about 20° C for 25-35 min; rinse; put into an ammino-silver solution (4.5% AgNO₃, 20 ml; pure ethanol, 10 ml; ammonia, sp. gr. 0.880, 2.4 ml; 2.5% NaOH, 1 ml) for 1-2 min; reduce in acidified formalin (distilled water, 400 ml; pure ethanol, 45 ml; 1 % citric acid, 13.5 ml; 10% formalin, 13.5 ml) for 1-3 min; wash; dehydrate through ascending grades of alcohol, including absolute; coat with 0.5% collodion, allow to dry slightly and harden in absolute alcohol-chloroform (2:1); rehydrate and put into 1% Na₂S₂O₃ for 1 min; dehydrate and cover.     

Counterstaining Golgi-Cox Impregnations with Luxol Fast Blue as a Myelin Stain

Immerse pieces of brain tissue 4 wk in solutions A and B, mixed just before use: A. K₂Cr₂O₇, 1 gm; HgCl₂, 1 gm; boiling distilled water, 85 ml. Boil A for 15 min, cool to 2 C and add: B. K₂CrO₄, 0.8 gm; Na₂WO₄, 0.5 gm; distilled water, 20 ml. Rinse in water and immerse 24 hr in LiOH, 0.5 gm; KNO₃, 15 gm; distilled water, 100 ml. Wash 24 hr in several changes of 0.2% acetic acid and then for 2 hr in tap water. Dehydrate and embed in celloidin. Process a 60 μ section through 70 and 95% ethanol, a 3:1 mixture of absolute ethanol and chloroform, and toluene. Immerse it for 5 min in a solution containing methyl benzoate, 25 ml; benzyl alcohol, 100 ml; chloroform, 75 ml. Orient the section on a chemically clean slide and let air-dry 5-10 min. Process through toluene, 3:1 ethanol-chloroform and 95% ethanol. Place the section for 5-60 min at 60 C in a solution made up of: Luxol fast blue G (Matheson, Coleman and Bell), 1 gm; 95% ethanol, 1000 ml; 10% acetic acid, 5 ml. Hydrate to water and immerse in 0.05% Li₂CO₃ for 3-4 min. Differentiate in 70% ethanol and place in water. Immerse for 5-15 min in a mixture of two solutions: A. cresylechtviolet (Otto C. Watzka, Montreal), 2 gm; 1 M acetic acid, 185 ml; B. 1 M sodium acetate, 15 ml; distilled water, 400 ml; absolute ethanol, 200 ml. Dehydrate to 3:1 ethanol-chloroform. Clear in toluene and apply a coverslip. The technique produces fast Golgi-Cox impregnated neurons against a background of counterstained myelinated fibers. Patterns of the myelinated fibers can be used to localize impregnated neurons.     

Gross Demonstration of the Mammalian Atrioventricular Bundle by A Periodic Acid-Schiff Procedure

The following procedure stains the atrioventricular conduction system selectively. (1) Wash the fresh heart with physiological saline solution to free it of blood; (2) fix it in 10% formalin containing 0.5% HIO₄ for 1 hr; (3) wash in 3 changes of distilled water for 20 min; (4) keep in 80% alcohol for 12 hr to 2 wk; (5) wash with distilled water; (6) treat with a dilute Schiff's reagent containing 0.1 gm of basic fuchsin per 100 ml for 0.5-2 min; (7) rinse in three changes of 2% Na₂SO₃ in 0.2 N HCI for 3-5 min; (8) wash and examine in 80% alcohol; store in 80% alcohol.     

Nile Blue Histochemical Method for Phospholipids

The technic recommended is: Fix 6-12 hr. in 10% formalin containing 1% CaCl₂. Cut frozen sections without embedding or after gelatin or carbowax. Stain 90 min. at 60°C. in saturated aqueous Nile blue sulfate, 500 ml. plus 50 ml. of 0.5% H₂SO₄, boiled 2 hr. before use. Rinse in distilled water, and place in acetone heated to 50°C. Remove the acetone from the source of heat and allow the sections to remain 30 min. Differentiate in 5% acetic acid 30 min., rinse in distilled water, and refine the differentiation in 0.5% HCl for 3 min. Wash in several changes of distilled water and mount in glycerol jelly. Results: phospholipids - blue; everything else - unstained. Counterstaining nuclei with safranin is optional, but if done, it preferably precedes the Nile blue and is then differentiated by the acetic acid. The histochemical principles on which the method is based are as follows: (1) The calcium compounds of phospholipids combine with the oxazine form of Nile blue sulfate and survive subsequent treatment; (2) neutral lipids are dissolved out by acetone; (3) proteins and other interfering substances are destained by the acetic acid and hydrochloric acid baths.     

Selective Staining of Neurosecretory Material in Semithin Epoxy Sections by Gomori's Aldehyde Fuchsin

The epoxy resin was removed from semithin (1 μm) sections by immersing them for 30 sec in sodium methoxide (Mayor et al., J. Biophys. Biochem. Cytol., 9: 909-10, 1961) and then processed as follows: (1) left for 1-3 hr at 60 C in a mixture of formalin, 25 ml; glacial acetic acid, 5 ml; CrO₃, 3 gm; and distilled water, 75 ml: (2) oxidized 10 min in a 1:1:6 v/v mixture of 2.5% KMnO₄, 5% H₂SO₄ and distilled water: (3) bleached in 1% oxalic acid, and (4) stained for 15 min in aldehyde fuchsin, 0.125% in 70% alcohol, or in a 1% aqueous solution of toluidine blue. The neurosecretory material is selectively stained.     

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.