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.     

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.     

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.     

The Quantitative Determination of Osmium Tetroxide in Fixatives

This rapid spectrophotometric method for determining the OsO₄ concentration in fixative and stock solutions is based on the reduction of OsO₄ by acidified KI to the blue species of OsI₆ =, which is then determined at 649 mµ. The salt K₂OsI₆ has been isolated from the reaction mixture and characterized. Method: A I ml aliquot of the solution, containing up to 3% OsO₄, is diluted to 100 ml with distilled water. To 1 ml of the diluted solution is added, in order: distilled water, 2 ml; 1 M HCI, 1 ml; and 1 M KI, 1 ml. Optical density at 649 mµ is read from 10-120 min thereafter. OsO₄ concentration is calculated from the measured molecular extinction coefficient of OsI₆ =, 4400 liter/mole cm.     

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).     

Methylene Violet (Bernthsen), by Zinc-Alkali-Chlorate Hydrolysis of Methylene Blue

Though Bernthsen's methylene violet (MV) is a common constituent of polychrome methylene blue, the hydrolytic oxydation of methylene blue to yield azure-free MV has been considered a difficult chemical reaction since the time of Bernthsen, who used Ag₂O in the hydrolysis. MV is qualitatively distinguished from azures by Bernthsen's criteria and the author's new tests: (1) light-excited isomeric change, (2) reactivity to acidity, (3) reaction with KCIO, and (4) reaction with Na₂SO₃ of azures in CHCI₃, while MV gives none. But MV shows (5) indicator properties at pH 4, while azures do not. For practical hydrolysis, treat methylene blue (10 parts by weight) and KCIO₃ (1 part) with 1-2 N NaOH to convert methylene blue to a mixture of MV and azures. Then dilute the solution, add a Zn salt and NaHCO₃ in excess of the amount needed to convert the NaOH to Na₂CO₃. Boil the solution gently for 1-2 hr. The end point of the reaction is found by pipetting a drop of reactant into 3% acetic acid in a test tube, adding CHC1₃ and extracting. The acetic layer should then be almost colorless while the CHC1₃ is colored intensely cherry red. After cooling, the precipitated dye is filtered and dried. This procedure gives good yields of a dye which meets the criteria given by Bernhsen. The peak of the absorption curve in solution, pH 4-11, is at 624 mμ (Bernthsen 625 mμ) and in acid solution, pH 0-4, 588 mμ (Biological Stains, 1953; 580 μ). The dye contains so little azures, that purification of the MV fraction obtained from the reaction mixture is unnecessary when it is used in the Wright-type Romanowsky stain. The remarkable staining effect of MV is its power to bring out red azurophil granules of monocytes and lymphocytes when used with eosinated thiazins in Wright's stain.     

Staining of Nerve Fibers with Methylene Blue Factors Improving the Staining

Several factors influencing the staining of nerve fibers with methylene blue, especially the influence of chloralhydrate and carbamylcholine chloride (as parasympathicotonics), and of some anesthetics were studied. The intestines of mouse, rat, and guinea pig were used. The following immersion technic is suggested: Tissue from animals anesthetised by chloralhydrate is immersed in: zinc free methylene blue, 0.03%; sodium tartrate, 0.5%; sodium pyruvate, 0.05% carbamylcholine, 0.00005%; 0.2 M Na₂HPO₄, 0.77%; 0.1 M citric acid, 0.18%; NACl, 0.79%; also an anesthetic which varies with the animal selected. Air is kept bubbling through the staining solution and microscopic examination is made at 6 min. intervals. After 0.5-1 hr. the tissue is fixed in: ammonium molyb-date, 10 g.; sucrose, 35 g.; distilled water, 100 ml.; to which is added just before use, 1% platinum chloride, 3 ml.; 2% osmic acid, 3 drops. Washing is in ice cold water and dehydration at 0°C. in Lang's fluids (varying mixtures of ethanol and n-butanol). The tissues thus prepared are stored in liquid paraffin.     

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.     

Bromphenol Blue as A Stain for Muscle Striations

Sections from formalin-fixed muscle are stained 4-24 hr with a 0.05% solution of bromphenol blue in 2% acetic acid, rinsed with water and placed in 0.5% acetic acid for 5-10 min. They are then treated 30 sec with tap water substitute (KHCO₃, 0.2 gm; MgSO₄, 2 gm; distilled water, 100 ml), rinsed, dehydrated in alcohol, cleared in xylene and covered in a polystyrene mountant Striatums of both cardiac and skeletal muscle fibers are fully resolved under oil immersion, against the blue background of the other parts of the fibers.     

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.     

Specific Staining of Sialic Acid Components on Sodium Dodecyl Sulfate Polyacrylamide Gel

Specific staining of sialic acid components after sodium dodecyl sulfate polyacrylamide gel electrophoresis can be carried out as follows: 1) extract glycoprotein of erythrocyte membranes or serum by the phenol-saline method, 2) electrophorese the extract on 5% polyacrylamide gel containing 0.1% sodium dodecyl sulfate at constant current, 3) treat the gel with chilled 0.04 M HIO₄ for 45 minutes, 4) replace the periodic acid solution with one containing resorcinol 0.6 g, cone. HCl 50 ml, 0.1 M CuSO₄ 0.5 ml and H₂O 50 ml, 5) warm the container in boiling water until blue violet sialic acid bands become clear, 6) replace the staining solution with a mixture of equal parts water and concentrated HCl and observe at once.     

Zinc Chloride Methylene Blue, Ii. Preparation of Giesma and Wright Type Low Azure B Blood Stains from Dichromate Oxidized Commercial Dye

TO determine the amount of K₂Cr₂O₇ required to produce optimal Giemsa type staining, six 1 g amounts (corrected for dye content) of zinc methylene blue were oxidized with graded quantities of K₂Cr₂O₇ to produce 4, 8, 12, 16, 20 and 24% conversion of methylene blue to azure B. These were heated with a blank control 15 minutes at 100 C in 60-65 ml 0.4 N HCI. cooled, and adjusted to 50 ml to give 20 mg original dye/ml. Aliquots were then diluted to 1% and stains were made by the “Wet Giemsa” technic (Lillie and Donaldson 1979) using 6 ml 1% polychrome methylene blue, 4 ml 1% cosin (corrected for dye content), 2 ml 0.1 M pH 6.3 phosphate buffer, 5 ml acetone, and 23 ml distilled water. The main is added last and methanol fixed blood films are stained immediately for 20-40 min.

For methylene blue supplied by MCB 12-H-29, optimal stains were obtained with preparations containing 20 and 24% conversion of methylene blue to azure B. With methylene blue supplied by Aldrich (080787), 16% conversion of methylene blue to azure B was optimal. Eosinates prepared from a low azure B/methylene blue preparation selected in this way give good stains when used as a Wright stain in 0.3% methanol solution. However, when the 600 mg eosinate solution in glycerol methanol is supplemented with 160 mg of the same azure B/methylene blue chloride the mixture fails to perform well. The HCI precipitation of the chloride apparently produces the zinc methylene blue chloride salt which is poorly soluble in alcohol. It appears necessary to have a zinc-free azure B/methylene blue chloride to supplement the probably zinc-free eosinate used in the Giemsa mixture.     

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.     

Nitric oxide stimulates prostaglandin synthesis in cultured rabbit gastric cells

Both prostaglandins (PGs) and nitric oxide (NO) have cytoprotective and hyperemic effects in the stomach. However, the effect of NO on PG synthesis in gastric mucosal cells is unclear. We examined whether sodium nitroprusside (SNP), a releaser of NO, stimulates PG synthesis in cultured rabbit gastric mucus-producing cells. These cells did not release NO themselves. Co-incubation with SNP (2 × 10⁻⁴, 5 × 10⁻⁴, 10⁻³ M) increased PGE₂ synthesis, and SNP (10⁻³ M) increased PGI₂ synthesis in these cells. Hemoglobin, a scavenger of NO, (10⁻⁵ M) eliminated the increase in PGE₂ synthesis by SNP, but methylene blue, an inhibitor of soluble guanylate cyclase, (5 × 10⁻⁵ M) did not affect the increase in PGE₂ synthesis by SNP. 8-bromo guanosine 3′ : 5′-cyclic monophosphate (8-bromo cGMP), a cGMP analogue, (10⁻⁶, 10⁻⁵, 10⁻⁴, 10⁻³ M) did not affect PGE₂ synthesis. These findings suggest that NO increased PGE₂ and PGI₂ synthesis via a cGMP-independent pathway in cultured rabbit gastric cells.     

Orthochromatic, Species-Limited Staining of Mast Cells With Night Blue

Night blue will stain the mast cells of rat, mouse and hamster selectively if alcohol differentiation is controlled. The technical steps are: Dewax paraffin sections with xylene, 2 changes; air dry; 2% Na₂SO₄, 3-5 sec; 0.5% night blue in 10% ethanol, 1 hr at 60°C; rinse in water; 9% HNO₃, 15 sec; water 1-5 min; 70% ethanol, 2 changes, 30 sec each; wash; 0.01% safranin, 3-5 sec; rinse, blot, air dry, mount in synthetic resin. A clear orthochromatic stain of the mast-cell granules occurs. Acid fixation prevents the staining reaction.     

Mercuric Bromphenol Blue Staining of Precipitin Lines in Agar

Lines formed by antibody-organ antigen reactions are stained particularly well by a modification utilizing the mercuric bromphenol blue (MBB) mixture of Mazia et al. (Biol. Bull., 104: 57-67, 1953). The agar covered slides are placed overnight in 0.85% NaCI at 4 C, followed by washing for 2 hr in 0.85% NaCI at 25 C. They are then rinsed for 10 min in distilled water, and dried overnight at 37 C. The precipitin lines are fixed by immersing the slides for 25 min in 95% alcohol, followed by 5 min hydration in distilled water. They are stained for 25 min in MBB mixture (HgCI₂, 10 gm; bromphenol blue, 0.1 gm; 95% ethanol, 100 ml). Excess stain is removed by immersing in acidified alcohol (95% ethanol, 98 ml; glacial acetic acid, 2 ml). Finally, the slides are passed through alcohol and xylene, and resin-mounted under coverslips.     

Lower Azure B Methylene Blue Ratios in Giemsa Type Blood and Malaria Stains

Starting from ancient reports that rare samples of methylene blue were apparently sufficiently contaminated with azures to give red plasmodial and red purple nuclear chromatin in Chenzinsky type methylene blue eosin stains, it was decided to determine how little azure B would suffice for such staining in methylene blue eosin stains. The traditional 1902 Giemsa had an azure:methylene blue: eosin ratio of about 6:3:6.3:10; Lillie's 1943 formula had a 5:7:10 ratio. In the current series of tests 5:7:10 (I), 4:8:10 (II), 3:9:10 (III), 2:10:10 (IV), 1:11:10 (V), and 0:12:10 (VI) were used. Malaria and blood stains were better than the standard 5:7:10 (I) in III, IV and II in that order. Normal and leukemic human blood, mouse blood with Plasmodium berghei, and monkey blood with the CDC strain of Pl. falciparum were used as test materials. The staining mixtures were made from highly purified samples of azure B and methylene blue. Staining mixtures contained 12 ml 0.1% thiazin dye, 10 ml 0.1% eosin, 2 ml each of glycerol, methanol and 0.1 M phosphate buffer pH 6.5, 3 ml acetone as accelerator, and distilled water to make 40 ml; staining times of 10-30 min were used.     

Studies on Polychrome Methylene Blue II. Acid Oxidation Methods of Polychroming

The action of K₂Cr₂O₇, Ag₂O, KMnO₄, HgO and NaIO₃ in polychroming methylene blue is explored. The last two have no action in neutral or acid methylene blue solutions. With the other three reagents the amount of polychroming, as measured by the shift in the absorption spectrum, is roughly proportional to the amount of oxidant used. Various lots of methylene blue produce similar products with similar proportions of K₂Cr₂O₇. With similar quantities of this reagent similar products are produced by polychroming at 100°, 80°, 70° or 60° C. At 100° C. the action of K₂Cr₂O₇ or of Ag₂O appears to be completed in 15 minutes. In K₂Cr₂O₇ polychroming, H₂SO₄ can be substituted for HCl, and subsequent BaCO₃ neutralization removes the salts formed and prevents accidental alkali polychroming. K₂Cr₂O₇ polychroming produces products with narrower absorption bands than alkali polychroming.     

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.     

Mordant Blue 3: a Readily Availablx Substitute for Hematoxylin in the Routine Hematoxylin and Eosin Stain

Mordant blue 3 may be used as a suhstitute for hematoxylin in hematoxylin and eosin stains. The staining solution consists of 0.25 g dye, 40 ml of 10% iron dam, 5 ml of cone H₂SO₄, and 955 ml of dirtilled H₂O. Staining the is 5 minutes, followed by differentiation in acid water or acid alcohol. After blueing, the seaions are counterstained with emin. Results closely resemble the hematoxylin and eosin stain.