Prestaining Oxidation by Acidified H2O2 for Revealing Schiff-Positive Sites in Epon-Embedded Sections

Reliable production and identification of Schiff-positive sites on glutaraldehyde-osmium fixed 0.5-1 μsm Epon sections is accomplished by preoxidation of sections with 10% H₂O₂ acidified with H₂SO₄ (HPSA) to pH 3.2 (Pool, C. R., Stain Techn., 44: 75-9, 1969). Light green as a counterstain is used. Steps in the procedure are: HPSA, 1-2 min at 25-30 C; washing; 1% light green 3-5 min; brief rinse; Schiff reagent 1-3 min; washing; drying; clearing in xylene and mounting in resin. The use of acidified H₂O₂ prevents the common occurrence of Schiff background staining in glutaraldehyde-fixed tissues and permits optimum penetration of staining solutions. Sections were attached to glass slides without adhesive and were processed in Coplin jars. Prior to drying, excess solutions should be drained and wiped away with lens tissue to prevent formation of precipitate on the sections.     

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.     

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).     

Silver Protein Staining of Nerve Fibers in Celloidin Sections

Celloidin sections from formalin-fixed brain and spinal cord of primates are stored in 70% alcohol after cutting, soaked in 2% pyridine in 50% alcohol for 6-8 hr at 37 C, and transferred to 1% concentrated NH₄OH in 50% alcohol 15-18 hr at 20-25 C. After washing and flattening, the sections are transferred to 1% silver protein solution containing 30 ml of 0.2 M H₃BO₃/100 ml. Impregnation is accomplished in 50 ml screw-top jars, 50 mm in diameter, which are filled to a depth of 35 mm, and have 1 gm of copper foil, 0.002 inch thick added. The foil is folded in loose accordion-fashion, pierced and threaded, cleaned in 5% HNO₃, rinsed in distilled water, and suspended in the solution just above the sections by fastening the thread to the jar lid. The sections are impregnated for 24 hr at 37 C, rinsed in distilled water, reduced in a solution of 5% Na₂SO₃ and 1% hydroquinone for 10 min, washed in distilled water and toned in 0.2% gold chloride for 5 min. After rinsing in distilled water, the sections are transferred to 1% oxalic acid for 45-60 sec, washed in distilled water and placed in 5% Na₂S₂O₃ for 5 min. Sections are then washed, dehydrated to 95% alcohol, cleared in terpineol, followed by 3 changes in xylene, and mounted.     

A Protargol Method for Staining Nerve Fibers in Frozen or Celloidin Sections

Extensive experimentation with protargol staining of neurons in celloidin and frozen sections of organs has resulted in the following technic: Fix tissue in 10% aqueous formalin. Cut celloidin sections IS to 25 μ, frozen sections 25 to 40 μ. Place sections for 24 hours in 50% alcohol to which 1% by volume of NH₄OH has been added. Transfer the sections directly into a 1% aqueous solution of protargol, containing 0.2 to 0.3 g. of electrolytic copper foil which has been coated with a 0.5% solution of celloidin, and allow to stand for 6 to 8 hours at 37° C. Caution: In this and the succeeding step the sections must not be allowed to come in contact with the copper. From aqueous protargol, place the sections for 24 to 48 hours at 37° C. directly into a pyridinated solution of alcoholic protargol (1.0% aqueous solution protargol, 50 ml.; 95% alcohol, 50 ml.; pyridine, 0.5 to 2.0 ml.), containing 0.2 to 0.3 g. of coated copper. Rinse briefly in 50% alcohol and reduce 10 min. in an alkaline hydroquinone reducer (H₃BO₃, 1.4 g.; Na₂SO₃, anhydrous, 2.0 g.; hydroquinone, 0.3 g.; distilled water, 85 cc; acetone, 15 ml.). Wash thoroly in water and tone for 10 min. in 0.2% aqueous gold chloride, acidified with acetic acid. Wash in distilled water and reduce for 1 to 3 min. in 2% aqueous oxalic acid. Quickly rinse in distilled water and treat the sections 3 to 5 min. with 5% aqueous Na₂S₂O₃+5H₂O. Wash in water and stain overnight in Einarson's gallocyanin. Wash thoroly in water and place in 5% aqueous phosphotungstic acid for 30 min. From phosphotungstic acid transfer directly to a dilution (stock solution, 20 ml.; distilled water, 30 ml.) of the following stock staining solution: anilin blue, 0.01 g.; fast green FCF, 0.5 g.; orange G, 2.0 g.; distilled water, 92.0 ml.; glacial acetic acid, 8 ml.) and stain for 1 hour. Differentiate with 70% and 95% alcohol; pass the sections thru butyl alcohol and cedar oil; mount.     

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.     

Staining of Oligodendroglia with Silver Ammino Tungstate in Unembedded and Embedded Material

Specimens of brain or spinal cord fixed in formalin, Cajal's formol-bromide, or Koenig, Groat and Windle's formalin-acacia can be used to stain oligodendrocytes in frozen, in paraffin, or in celloidin sections. The sections are soaked 3-5 min in 0.02% acetic acid, pH 3.4, then rinsed 2-3 sec in 3% H₂O₂ and transferred to a silver bath prepared as follows: Mix equal parts of 10% AgNO₃ and 10% Na₂WO₄, and dissolve the precipitate with concentrated NH₄OH; avoid an excess of ammonia. Silver at room temperature for 15-20 sec, develop in 1% formalin, dehydrate, and mount. For embedded material, prepare a mixture consisting of 1 part of 10% aqueous Aerosol MA and 4 parts of 10% Aerosol OT in 95% alcohol. Add 5 drops of this mixture to each 50 ml of dilute acetic acid and 3% H₂O₂; 5 drops to each 20 ml of the silver bath.     

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.     

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.     

Localization of Myoglobin in Striated Muscle of the Domestic Pig; Benzidine and NADH2-TR Reactions

Tissue blocks 1 cm³ from longissimus (white) and trapezius (red) muscles of adult pigs were fixed in phosphate-buffered 2.5% glutaraldehyde, pH 7.4, for 4 hr at about 25 C; washed 4 hr in running tap water, and immersed in 30% w/v sucrose solution for 16 hr or more. After freezing in liquid N₂, cryostat sections were cut and floated into saturated aqueous benzidine containing 0.15% H₂O₂ at 25 C for 30 min. Stained sections were washed in distilled water and mounted on slides with glycerol jelly. Three distinguishable gradiations of color intensity were found: strong, intermediate, and negative. The trapezius had a greater number of myoglobin-positive fibers than the longissimus muscle. Myoglobin-positive and myoglobin-negative staining occurred in red and white fibers, respectively; intermediates were apparently more closely related to the red than to the white fibers. The NADH₂TR reaction showed the same sites as did the benzidine reaction.     

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.     

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.     

Critical Staining of Pancreatic Alpha Granules with Phosphotungstic Acid Hematoxylin

Mammalian pancreatic alpha granules were differentially stained with phosphotungstic acid haematoxylin. Paraffin sections were dewaxed and hydrated, oxidised 5-40 sec in freshly prepared 0.3% KMnO₄ acidified with 0.3% (w/v) H₂SO₄, decolourised in 4% potassium metabisulphite, mordanted 20 min to 2 hr in 4% iron alum, stained in phosphotungstic acid haematoxylin 16-48 hr, rinsed in 95% ethanol until no stain runs from the tissue, dehydrated in absolute ethanol, cleared in xylene, and covered in synthetic resin. Advantages of this procedure are: (1) consistent, reproducible staining; (2) applicability to all the common laboratory mammals and man; (3) wide latitude at each stage, permitting its use as a routine method; and (4) superior visualization of alpha granules, due to suppression of background staining and absence of glare. For fixation, formalin-acetic or Bouin's solution is recommended.     

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.     

Salivary hydrogen peroxide produced by holding or chewing green tea in the oral cavity

Tea (Camellia sinensis) catechins have been studied for disease prevention. These compounds undergo oxidation and produce H₂O₂. We have previously shown that holding tea solution or chewing tea leaves generates high salivary catechin levels. Herein, we examined the generation of H₂O₂ in the oral cavity by green tea solution or leaves. Human volunteers holding green tea solution (0.1-0.6%) developed salivary H₂O₂ with C_max = 2.9-9.6 μM and AUC_0 → ∞ = 8.5-285.3 μM min. Chewing 2 g green tea leaves produced higher levels of H₂O₂ (C_max = 31.2 μM, AUC_0 → ∞ = 1290.9 μM min). Salivary H₂O₂ correlated with catechin levels and with predicted levels of H₂O₂ (C_{max(expected)} = 36 μM vs C_{max(determined)} = 31.2 μM). Salivary H₂O₂ and catechin concentrations were similar to those that are biologically active in vitro. Catechin-generated H₂O₂ may, therefore, have a role in disease prevention by green tea.     

Staining Tissue of the Central Nervous System with Luxol Fast Blue and Neutral Red

The tissue is fixed in 10% neutral saline formalin for 1 day to 3 wk depending on the size of the block, dehydrated and embedded in paraffin. The sections are stained at 57° C for 2 hr, then at 22° C for 30 min, in a 0.0125% solution of Luxol fast blue in 95% alcohol acidified by 0.1% acetic acid. They are differentiated in a solution consisting of: Li₂CO₃, 5.0 gm; LiOH-H₂O, 0.01 gm; and distilled water, 1 liter at 0-1° C, followed by 70% alcohol, and then treated with 0.2% NaHSO₃. They are soaked 1 min in an acetic acid-sodium acetate buffer 0.1 N, pH 5.6, then stained with 0.03% buffered aqueous neutral red. Sections are washed in distilled water, 1 sec, then treated with the following solution: CuSO₄·5H₂O, 0.5 gm; CrK(SO₄)₂·12H₂O, 0.5 gm; 10% acetic acid, 3 ml; and distilled water, 250 ml. Dehydration, clearing and covering complete the process. Myelin sheaths are stained bright blue; meninges and the adventitia of blood vessels are blue; red blood cells are green. Nissl material is stained brilliant red; axon hillocks, axis cylinders, ependyma, nuclei and some cytoplasm of neuroglia, media and endothelium of blood vessels are pink.     

Controlled Formaldehyde-Catecholamine Condensation in Cryostat Sections to Show Adrenergic Nerves by Fluorescence

Three sets of sections of freshly removed tissue are cut at 18 μ in a cryostat and dried on slides for 1.5 hr over P₂O₅. Each set of sections is incubated with a differently hydrated paraformaldehyde (prepared by storing paraformaldehyde powder over 21%, 25% or 28% aqueous H₂SO₄ for 1 wk) at 80 C for 1 hr before being mounted in glycerol and viewed with a fluorescence microscope. At least one set of specimens shows optimal fluorescence. The entire procedure from removing the tissue to observing fluorescence microscopically is accomplished readily within 4-8 hr. Adrenergic axons in the medial muscle of the cat nictitating membrane, the myometrium of the cat uterus and the adventitia of arterial vessels in rat pancreas are demonstrated.     

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.     

Hydrogen peroxide degradation by immobilized cells of alkaliphilic Bacillus halodurans

Whole cells of Bacillus halodurans LBK 261 were used as a source of catalase for degradation of hydrogen peroxide. The organism, B. halodurans grown at 55°C and pH 10, yielded a maximum catalase activity of 275 U g^-1 (wet wt.) cells. The catalase in the whole cells was active over a broad range of pH with a maximum at pH 8-9. The enzyme was optimally active at 55°C, but had low stability above 40°C. The whole cell biocatalyst exhibited a K_m of 6.6 mM for H₂O₂ and V_max of 707 mM H₂O₂ min^-1 g^-1 wet wt. cells, and showed saturation kinetics at 50 mM H₂O₂. The cells were entrapped in calcium alginate and used for H₂O₂ degradation at pH 9 in batch and continuous mode. In the batch process, the immobilized preparation containing 1.5 g (wet wt.) cells could be recycled at least four times for complete degradation of the peroxide in 50 mL solution at 25°C. An excess of immobilized biocatalyst could be used in a continuous stirred tank reactor for an average of 9 days at temperatures upto 55°C, and in a packed bed reactor (PBR) for 5 days before the beads started to deform.     

Pseudoisocyanin Staining of Insulin and Specificity of Empirical Islet Cell Stains

Two closely related pseudoisocyanins, N,N'-diethyl-6,6'-dichlorpseudoisocyanin chloride and N, N'-diethylpseudoisocyanin chloride, were tested for their metachromatic staining behavior with oxidized insulin. N,N'-diethyl-6,6-dichlorpseudoisocyanin chloride gave nonspecific metachromasia with collagen, mucus, and mast cells of adult tissues; almost all tissues of rat embryos exhibited nonspecific staining. Nonspecific reactions were rarely observed in adult or fetal tissues with the extremely labile metachromasia of N, N'-diethylpseudoiso-cyanin chloride. When oxidation time and temperatures are carefully controlled, this reagent apears to be highly specific for insulin-containing cells and can be used as a selective stain for beta cells. Paraffin sections of formalin fixed material were oxidized 45 sec at 28-29 C in freshly prepared acidified permanganic (2.5% KMnO₄, 1; 5% H₂SO₄, 1; distilled water, 7—parts by volume), decolorized 30 sec in 5% oxalic acid, and washed 5 min in running tap water. After rinsing in 2 changes of distilled water, sections were stained 20 min in a 36 mg/100 ml aqueous solution of N, N'-diethylpseudoisocyanin chloride. Sections were then washed in running tap water until the albumen adhesive was decolorized, and mounted in Karo syrup diluted with an equal amount of distilled water. The insulin-containing cells are stained light to dark purple; all other tissue components, various shades of red. N, N'-diethylpseudoisocyanin chloride was used as a reference for evaluating the specificity of 5 commonly used empirical methods for demonstrating alpha and beta cells in pancreatic islets. Cells exhibiting pseudo isocyanin metachromasia were stained selectively by aldehyde-fuchsin, Heidenhain's azan, and chrome-hematoxylin. Aldehyde-Iuchsin was the only empirical stain tested which gave results comparable to pseudoisocyanin for clarity and definition of beta cells. After oxidation in acidified permanganate, azocarmine and phosphotungstic acid-hematoxylin differentially stained alpha cells; cells demonstrated by these two methods did not exhibit pseudoisocyanin metachromasia. This histochemical procedure can precede empirical methods which require preliminary oxidation in acidified permanganate or it can follow empirical methods which do not extract the insulin nor alter its intramolecular disulfide bonds.