A Simple Histochemical Reaction for Aldehydes

A study has been made on the possibility of replacing leucofuchsin by colored basic fuchsin for the histochemical demonstration of aldehydes. Several tissues from mammals and various pertinent fixatives were used. Aldehydes were freed from carbohydrates by oxidation and from thymonucleic acid by hydrolysis.

It was found that the colored form and not necessarily the leucoform of basic fuchsin can be used histochemically in demonstrating aldehydes. The technic used is as follows: (1) Treat with 1.0-0.5% H₅IO₆ (or in 1% KIO₄ in M/1 H₂SO₄) for 5 to 10 min. and wash thoroughly. For thymonucleic acid hydrolize with N HCl 5 min. at room temperature, 10 min. at 60°C. and 5 min. at room temperature. (2) Stain for 2-3 min. with 0.05% basic fuchsin in 5% ethanol, 3% phenol. (3). Transfer immediately to 1 or 2 changes of 1% sodium bisulphite or potassium metabisulphite in 0.1-0.2 N H₂SO₄ for a total of 5 min. (4) Rinse with water and treat with M H₂SO₄ in 95% ethanol for 3-5 min. 6. Wash thoroughly in water and dehydrate, clear, and mount. For glycogen and mucin the following counterstaining solution is recommended: orange G, 0.25 g.; light green SFY, 0.10 g.; phosphotungstic acid 0.50 g.; 50% ethanol, 100 ml.; glacial acetic acid, 0.25 ml.     

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).     

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.     

Differential in Toto Staining of Bone, Cartilage and Soft Tissues

The following method for staining bone and cartilage allows study of the gross cleared specimen and does not injure the tissues for subsequent microscopic study: Fix in 10% neutral formalin; bleach thoroughly in 3% H₂O₂ in sunlight. Wash in distilled water. Stain bone 24 hours in 0.01 g. of Biebrich scarlet in 100 ml. of distilled water. Destain in 95% alcohol until soft tissues and cartilage are colorless. Stain cartilage 24 hours in a pH2 buffer solution of 2.1g. of citric acid per 100 ml. of water with 0.001 g. of methylene blue. Destain in pH2 buffer solution until soft tissues are pale. Dehydrate in two changes of 95% alcohol in preparation for clearing. (This completes the destaining and may remove too much stain from the cartilage if destaining in the pH2 solution has been carried too far.) Place in Groat's clearing fluid and cover loosely so that the alcohol may evaporate, or remove the alcohol in vacuo. Groat's Mixture No. 19 is usually satisfactory.

For a combined stain, first stain bone, as above, and then apply the cartilage stain.

Seal jars with an ordinary liquid wood glue such as LePage's.     

Haematoxylin-Phloxine-Alcian Blue-Orange G Differential Staining of Prekeratin, Keratin and Mucin

This is a modification of Kreyberg's stain with Alcian blue 8GS used to stain acid much while phloxine B and orange G stain keratin and prekeratin. Procedure: Dewax formalin-fixed paraffin sections in xylene and hydrate through alcohol. Stain in Mayer's haemalum, 10 min; blue in tap water; wash in distilled water; stain in 1% phloxine, 3 min; wash in running water, 1 min; wash in distilled water; stain in 0.5% aqueous Alcian blue in 0.5 acetic acid, 5 min; wash in distilled water; stain in 0.5% orange G dissolved in 2.0% phosphotungstic acid, 13 min; dehydrate quickly in 2 changes of 95% alcohol and 2 changes of absolute alcohol; clear in several changes of xylene; mount in a synthetic resin. Acid mucopolysaccharides are stained turquois blue; prekeratin and keratin are orange to red orange.     

Feulgen's Reaction Applied to Protozoa and Small Worms, Mounted In Toto in Venetian Turpentine

Permanent mounts of certain protozoa and small worms are obtained as follows: kill suspensions of the organisms with Feulgen's fixative (6% HgCl₂ in 2% aqu. acetic acid) for 3 to 24 hours. After pipetting off the fixative, add successively: 70% iodized alcohol; ditto, 30 minutes later; 50%, then 35% alcohol; 2 baths distilled water; normal HCl. Transfer to cold water and heat to 60°C for 4 to 5 minutes or longer. Cool under running water; and wash in distilled water.

Stain 1 to 3 hours in Feulgen's fuchsin sulfurous acid (1 g. of a suitable basic fuchsin, e. g. rosanilin hydrochloride, boiled in 200 cc. water, cooled, and allowed to stand 24 hours after adding 20 cc. normal HCl and 1 g. sodium bisulfite). Pass thru 3 baths of 200 cc. distilled water with 10 cc. normal HCl and 1 g. sodium bisulfite. Transfer to water and then to 35%, 70%, and 95% alcohols successively. Counterstain with fast green FCF, orange G or eosin Y in 95% alcohol. Pass thru two changes of absolute alcohol.

Transfer to 10% Venetian turpentine and place in a dessicator; mount after the turpentine has become concentrated.

If sections instead of total mounts are desired, the material should go from absolute alcohol, thru alcohol-xylol and xylol to paraffin (or preferably paraffin of M. P. 56°C with 3% bees-wax). The paraffin may be added to the material in the test tube, and cooled after the organisms have settled. Then break the tube, trim a block, and cut.     

Glutaraldehyde-Ammoniacal Silver Carbonate-Formaldehyde Staining of Histones of Mitotic Chromosomes

Cells from monolayer culture of Chinese hamster line Don were treated by Colcemid (0.1 μg/ml) for 2 hr, trypsinized and spun; resuspended in 0.5% sodium citrate solution for 10 min, respun, and then resuspended in a small volume of the supernatant. Slide preparations were made by smearing, followed by air drying for 1 min at room temperature. They were fixed and stained by the following sequence: 2.5% glutaraldehyde in Millonig's buffer, 30 min; distilled water, 6 min, 5 changes; ammoniacal silver at 18-26 C, 10 sec; distilled water, 30 min, 5 changes; 2.5% formalin, 2 min; and distilled water, 3 changes during 15 min. Staining solution: add 225 ml of 5% Na₂CO₃ to 75 ml of 10% AgNO₃, then add concentrated NH₄OH slowly, drop by drop, until the solution is transparent. Finally add 300 ml of dstilled water. Cells treated with cold 0.25 N HCl before fixation were not stained. Sequence modifications show that chromatin does not reduce silver by itself. This method stains the sites of high histone concentrations in mitotic chromosomes of cytogenetic preparations.     

Acid Orcein-Iron and Acid Orcein-Copper Stains for Elastin

Paraffin sections of formol-fixed tissues stained 4-18 hr in 70% alcohol containing 1% orcein and 1% of concentrated (12 N) HCl by volume yield the familiar purple brown elastin and red nuclei on a pink background. When sections so stained are transferred directly from the stain to 70% alcohol containing 0.02% ferric chloride (FeCl₃·6 H₂O) or 0.02% copper sulfate (CuSO₄·5 H₂O) for a 15 sec to 3 min period, elastin coloration is changed to black or reddish black and chromatin staining to reddish black. The procedure can be counterstained with picro-methyl blue to yield blue collagen and reticulum or with our flavianic acid, ferric chloride, acid fuchsin mixture to give deep yellow background and deep red collagen.     

Staining of fresh, Undecalcified, thin Bone Sections

Fresh, undecalcified bone sections can be reproducibly and reliably stained by any of the following procedures: (A) Basic fuchsin, 1% in 30% alcohol, 48 hr, 22°C. (B) AgNO₃, 0.033 M, 48 hr, 22°C; washing 48 hr in a large volume of distilled water; exposure to light to develop the color. (C) Metallic sulfides (Co⁺⁺, Pb⁺⁺, Hg⁺⁺, Cu⁺⁺): the nitrate of the metal, 0.033 M, 48 hr, 22°C; then Na₂S, 0.033 M, 48 hr, 22° C. (D) Alizarin Red S, 0.1% solution in distilled water, 48 hr, 22°C; differentiated 48 hr at 22°C in weakly alkaline water, pH about 8. (E) KMnO₄: boiling 8-10 min in a 0.1 N, solution. With the exception of D the surface stain must be ground off the section for microscopic examination of its interior. Stain concentration, time and temperature can be altered to suit specific needs.     

Histochemical Demonstration of Pyrophosphatase

Fresh tissue slices were fixed in 5% formalin containing 0.9% NaCl for 10-20 min and frozen sections therefrom floated for 3 hr at 37°C on an incubating mixture made as follows. Sodium pyrophosphate (Na₄P₂O₇-12H₂O), 1.088 gm was dissolved in 20-30 ml of distilled water and to this was added ferric chloride (FeCl₃-6H₂O), 0.61 gm dissolved in 10-15 ml of water. The precipitate was just dissolved by cautiously adding 5-10% aqueous Na₂CO₃ solution and the pH adjusted to 7.2 with 1N HCl. The volume was made up to 100 ml and 0.9 gm of NaCl added. Before use, 1 ml of 10% Mg(NO₃) was added. After incubation, sections were washed 10-15 min in 0.9% NaCl, then mounted on glass slides and air-dried. When dry, the slides were immersed in 0.9% NaCl containing 0.2-0.5% ammonium sulfide for 2-3 min, then dehydrated rapidly through graded alcohols, cleared, and covered in balsam. Sites of pyrophosphatase activity stained in various shades of green. Acid pyrophosphatase also was histochemically demonstrated by the same principle, excepting that the substrate solution was adjusted to pH 3.7-4.0 with acetate buffer. The pattern of distribution of pyrophosphatase and glycerophosphatase was almost identical.     

A Method for the Selective Removal of Deoxyribonucleic Acid from Tissue Sections

The deoxyrihonucleic acid (DNA) of chromatin undergoar depurinization on mild acid hydrolysis with a picric acid-formaldehyde mixture (Bouin's fluid). The apurinic acid thus formed is degraded by condensation with aniline and is lost from tissue sections, but ribonucleic acid (RNA) in nucleoli and cytoplasm is well preserved. Technique: Fi in Carnoy's fluid (ethanol:acetic acid 3:1 or ethanol:chloroform:acetic acid 6:3:1) or in aldehydes (10% formalin or 2.5% glutaraldehyde bsered to pH 7.0). Hydrolyse deparaEnii sections 12-24 hr at 27-50 C in Bouin's fluid, wash in distilled water, immerse in 25% (v/v) acetic acid, treat 1 hr at 27-30 C with 10% (v/v) dine in 25% acetic acid, wash in 25% acetic acid and then in water. Stain 10-40 min with 0₃% toluidine blue in 0.05 M potassium biphthalate bder (pH 4.0); rinse in distilled water, pass to 10% (w/v) ammonium molybdate for 1 min, rinse again in water and pass through tert-butanol and xylene to a synthetic resin. Chromatin and chromosomes are pale green; RNA in nucleoli and cytoplasm deep purple.     

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.     

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 and Mounting Helminths

The following procedure is recommended: Fix ces-todes and trematodes (while held flat between glass slides) 0.5-2.0 hr. in the following mixture: formalin, 15; acetic acid (gl.), 5; glycerol, 10; 95% ethyl alcohol, 24; distilled H₂O, 46; all proportions by volume. After freeing them from the slides, wash thoroughly in running water and stain immediately thereafter. Stock staining solution: ferric ammonium alum (violet cryst.), 2 g.; distilled H₂O (cold) 100 ml.; after solution, add 2 ml. concentrated H₂SO₄, bring to a boil; add 1 g. coelestin blue B (Nat. Aniline), boil 3-5 min.; cool and add 10 ml. absolute methyl alcohol and 10 ml. glycerol. Dilute 1 vol. with 3 vol. distilled H20 for use. Stain 5-30 min., depending on size of specimens. Wash with 2 changes 0.5 hr. each of distilled H₂O, then 50% isopropyl alcohol 12-16 hr., 50% isopropyl alcohol 2 hr., followed by graded isopropyl alcohol for dehydration. Ether: ethyl alcohol (equal parts), 1 hr., is followed by embedding in celloidin in a sheet just thick enough to cover the specimens. Trim embedded specimens and dehydrate with isopropyl alcohol, 80%, 90% and absolute. Clear in beechwood creosote. Mount in balsam with cover glasses that overlap the edges of the celloidin 1-2 mm. While drying at 37°C, refill edges of mount with fresh balsam as needed. When dry, remove excess balsam and ring the edges with ordinary gloss enamel paint.     

Studies on Textile Dyes for Biological Staining II. A Trichrome Stain for General Use

This trichrome staining procedure differentially stains elastic fibers, collagen fibers and mucin. Gomori's aldehyde-fuchsin is used for elastic fibers; fast yellow TN is the component used for collagen and cytoplasm; pontacyl blue black SX is the nuclear stain. Procedure: Paraffin sections to water; aldehyde-fuchsin, 30 min; 70% ethanol; distilled water; 0.75% pontacyl blue black SX in 1.5% K.₂Cr₂O₇, 15 min; tap water; 70% ethanol to wash off all free dye; 2% fast yellow TN in 95% ethanol, 5 min; dehydrate, clear and cover.     

Enhanced Differentiation of Zaprionus Chromosomes by Prestaining Acid Hydrolysis

Two variations of orcein staining have been adapted to salivary gland chromosomes of Zaprionus. Method I: after dissection, glands are transferred to 1 N HCl at 60° C for 5 min, stained with 2.5% orcein in 60% acetic add for 15-20 min, and squashed in 60% acetic acid. Method II: after dissection, glands are transferred to 1 N HCl at 60° C for 5 min, transferred to a saturated solution of carmine in 45% acetic acid for 1 min, then to a mixture of 50 ml of 1% orcein in concentrated lactic acid and 50 ml of 30% acetic add for 5 min. They are squashed in the same mixture. The unproved differentiation of chromosomes from cytoplasm is attributed to the removal of cytoplasmic ribonucleic add by add hydrolysis.     

Handling Germinated Pollen on Millipore Membrane During the Feulgen and Autoradiographic Procedures

To prevent loss of pollen during the Feulgen's procedure, the pollen was grown on an autoclaved membrane filter (Millipore AA WP 025 00) in contact with a sterilized medium containing agar 0.5-1%, sucrose according to the genus (Malus 0.3-0.5 M; Persica and Tulipa 0.4 M), and H₃BO₃, 0.01%. To fix the germinated pollen of most species, the membrane was placed for 2 hr to overnight at 2-4 C on filter paper wet with the following mixture: OsO₄, 1 gm; CrO₃, 1.66 gm; and distilled water, 233 ml. To fix Persica pollen, 10% of glacial acetic acid had to be added to the fixative. Washing with distilled water and bleaching with a mixture of 3% H₂O₂ and sat. aq. ammonium oxalate, 1:1, were performed also on filter paper. Similarly, the preparation was processed for Feulgen staining by use of pieces of filter paper wet with the required fluids. Hydrolysis preceding the Schiff's reagent was performed at room temperature with 5 N HCl for 18 min. The differentiation after the Schiff's action was with 2% K₂S₂O₅ buffered to pH 2.3 with 9 ml of phosphate buffer (KH₂PO₄, 1.4 gm; conc. HCl, 0.35 ml and distilled water to make 100 ml). The stained pollen was floated off the membrane with a drop of glacial acetic acid to a gelatinized or an albumenized slide, and squashed. When the coverslip is removed the preparation may be either dehydrated and mounted or coated with autoradiographic film.     

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.     

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.     

The Diazosafranin Method; Control of Nitrite Concentration and Refinements in Specificity

Safranin is diazotized by using the customary molar ratio—dye, 1:HCl, 3:NaNO₂l. Partly oxidized NaNO₂ can be used, if necessary, by increasing the concentration of its solution enough to cause the normal color change from red to deep blue to occur within 2 min after adding the NaNO₂. To avoid carrying over excess HNO₂ into the alkaline coupling solution, 1 ml of 3% alcoholic urea solution (30 mg) is added for each milliliter excess of 1 N NaNO₂ used. Any free HNO₂ remaining at the end of the diazotization period produces a deep blue violet on starch-KI paper. Prolonged acid washing may be applied after coupling to decolorize cationic dye staining or triazenes. Na₂S₂O₄, TiCl₃ or SnCl₂ may be used to bleach true azo colorations. This decolorization is not limited to newly formed azo compounds with tissue.