Azophloxine in Two-Dye and Four-Dye Staining Combination

Tissues are fixed in either 10% formalin or Lavdow-sky's mixture. After the tissues are sectioned and mounted, they are stained in hematoxylin, then counterstained for 2 minutes in 0.1% aqueous azophloxine to which 4 drops of acetic acid have been added to each 100 ml. of stain. Sections are then rinsed in 0.2% acetic acid and dehydrated. Azophloxine GA can be used also in a tetrachrome method. Sections are stained in Harris' hematoxylin, washed, and placed in 0.2% acidified aqueous azophloxine for 2 minutes. They are then rinsed in 0.2% acetic acid, stained 1 minute in an aqueous mixture of 4% phosphotungstic acid and 2% orange G solution and rinsed again in 0.2% acetic acid. Finally, they are stained in 0.2% light green for 2 minutes, and differentiated in 0.2% acetic acid for 5 minutes. The advantage in using azophloxine is that it is clear and delicate and when used in a constant concentration, does not overstain if the recommended procedure is followed.     

Modified Whipf's Polychrome: A Connective Tissue Stain with Special Application for Demonstrating Leishmania

A polychrome stain procedure was developed to demonstrate amastigotes of the protozoan parasite Leishmania braziliensis as well as cytoplasmic and other tissue components in cutaneous lesions of infected animals. The procedure is as follows: stain nuclei for 10 minutes with an iron hematoxylin containing 0.5% hematoxylin and 0.75% ferric ammonium sulfate dissolved in 1:1 0.6 N H₂SO₄:95% ethanol; rinse 4 minutes in distilled water. Cytoplasmic staining is achieved by exposing tissues for 10 minutes to a solution containing 0.25% Biebrich scarlet, 0.45% orange G, 0.5% phosphomolybdic acid and 0.5% phosphotungstic acid in 1% aqueous acetic acid. These first two solutions are modified from Whipf's polychrome stain. Sections are differentiated for 10 seconds in 50% ethanol, rinsed in water, stained 3 minutes in 0.1% aniline blue WS in saturated aqueous picric acid, rinsed in water and differentiated for 1 minute in absolute ethanol containing 0.05% acetic acid. Mordanting overnight in 6% picric acid in 95% ethanol produced optimal results.

This procedure was applied to sectioned material from experimental animals with various protozoa. Trypanosoma cruzi, Besnoitia Jellisoni, Toxoplasma gondii and especially Leishmania braziliensis were well demonstrated. Combining cytoplasmic dyes and phosphomolybdic-phosphotungstic acids into one solution afforded differential staining of tissues by Biebrich scarlet and orange G; connective tissues were stained by this solution. Substantially improved definition of connective tissues resulted after counterstaining. This procedure differs from the Massou sequence in which connective tissues are first stained by cytoplasmic dyes along with other tissues and then destained prior to specific counter-staining. in comparing dyes structurally related to Biebrich scarlet, it was found that Crocein scarlet MOO, but not Poncenu S, was an acceptable substitute. Sirius supra blue GL and Sirius red FSBA were not useful as replacements for aniline blue WS in this procedure.     

Azophloxine in Two-Dye and Four-Dye Staining Combination

Tissues are fixed in either 10% formalin or Lavdow-sky's mixture. After the tissues are sectioned and mounted, they are stained in hematoxylin, then counterstained for 2 minutes in 0.1% aqueous azophloxine to which 4 drops of acetic acid have been added to each 100 ml. of stain. Sections are then rinsed in 0.2% acetic acid and dehydrated. Azophloxine GA can be used also in a tetrachrome method. Sections are stained in Harris' hematoxylin, washed, and placed in 0.2% acidified aqueous azophloxine for 2 minutes. They are then rinsed in 0.2% acetic acid, stained 1 minute in an aqueous mixture of 4% phosphotungstic acid and 2% orange G solution and rinsed again in 0.2% acetic acid. Finally, they are stained in 0.2% light green for 2 minutes, and differentiated in 0.2% acetic acid for 5 minutes. The advantage in using azophloxine is that it is clear and delicate and when used in a constant concentration, does not overstain if the recommended procedure is followed.     

The Germination and Staining of Basidia in Gymnosporangium

A procedure is described for germinating and staining rust teliospores on the slide. The spores are germinated on slides in a damp chamber, about 3 hours being required for the production of sporidia. The material is killed by inverting slides over osmic acid fumes for a few minutes. Germinated spores are then allowed to dry on the slide, thus becoming fixed to the slide in a gelatin produced by the breaking down of their own stalks during germination. No other fixative is required. Material must be thoroly dehydrated in the alcohols (one or more hours in each of the higher alcohols); returned to water; mordanted for 2-3 hours in 4% iron alum; stained for 2-3 hours in 0.5% aqueous solution of Heidenhain's hematoxylin; destained in 2% iron alum. The material is passed back thru the alcohols and mixtures of xylol and absolute alcohol (1:2, 1:1, 2:1) to xylol and mounted in balsam. The method is particularly satisfactory for the Gymnosporanghim rusts, which have telia very readily gelatinized. The details of germination are preserved intact, as in nature, and many details of nuclear division are excellent.     

Gram Reaction in Crushed Yeasts

Thick smears of yeasts are dried, crushed between slides or cover-slips, stained with anilin crystal violet 11/2 minute, Lugol's iodine solution 2 minutes, decolorized in undiluted anilin oil, and counterstained with 1% aqueous safranin. Photomicrographs are given of crushed yeasts, thus stained, which show a Gram-positive outer layer and a Gram-negative inner substance.     

Differential Staining with a Mixture of Safranin and Fast Green FCF

Mounted deparaffinized sections were stained for 30-60 minutes at room temperature in a mixture of equal volumes of 0.1% aqueous solutions of safranin O and fast green FCF filtered before use. They were then washed in distilled water for 5 minutes, blotted, washed in 2 changes of absolute alcohol (2-3 min) and mounted from xylene. The nucleic acids are stained purplish-red, half esters of sulfuric acid orange, and proteins green. The procedure is applicable to a variety of materials fixed in a number of reagents though best results are obtained after acetic-alcohol fixation. Bouin's fluid and 10% neutral formalin are not suitable fixatives for this procedure. After acetic-alcohol fixation, the staining procedure may be used in conjunction with enzyme or extraction technics in order to characterize certain chemical components of cells or tissues. The safranin-fast-green technic has proved useful in investigations of pathological changes in tissues; in the visualization of secretory granules and in studies of cellular differentiation. The technic also would appear to greatly facilitate mitotic index determinations.     

Suggested Counterstains for Davenport Reduced Silver Preparations of Peripheral Nerves

—Peripheral nerves which have been fixed in a mixture of formaldehyde and acetic acid and stained according to the method of Davenport can be successfully counterstained for demonstration of myelin sheaths and stroma. After mounted sections have been silvered, reduced and toned, the coating of nitrocellulose is removed by passing thru two changes of acetone. Following brief washes in 100,95,85 and 75% alcohols they are stained in an acidified aqueous solution of azo carmine for 30 to 60 minutes. Excess azo carmine is extracted with anilin alcohol followed by acetic alcohol after which the sections are mordanted for 15 to 60 minutes in a 5% aqueous solution of phosphotungstic acid. Without washing they are transferred to a stain mixture of either anilin blue and orange G (acidified) or light green and orange G (acidified) where they remain from 1 to 5 hours. After destaining in 95% alcohol and dehydration in absolute alcohol the sections are mounted in dammar. Result: axons stain black; sheath and fibroblast nuclei, red; myelin sheaths, orange; and connective tissue, blue or green. When the counterstains are applied to ganglia, cytological details of individual cells are demonstrated.     

Mallory's Connective Tissue Stain Following Hematoxylin

The following combination of hematoxylin with Mallory's connective tissue stain is useful in bringing out nuclei as well as in differentiating tissue:

Slightly overstain in Mayer's hematoxylin (50 g. potassium alum and 0.2 g. sodium iodate added to 1 liter 0.1% aqueous hematoxylin). Wash; and stain 30 seconds to 1 minute in 0.04% aqueous acid fuchsin-Stain 4 minutes in: 0.5 g. anilin blue and 2 g. orange G dissolved hi 100 cc. of 1% aqueous phosphomolybdic acid. Pass thru 95% alcohol to absolute; clear in xylol and mount in balsam.     

A Differential Staining Method for Elastic Fibers, Collagenic Fibers, and Keratin

A method allowing for the differential presentation of elastic fibers, other connective tissue fibers, epithelial and other types of cytoplasm, and keratin is described. The procedure is based on the affinity of orcein for elastic fibers, of anilin blue for collagenic material, and of orange G for keratin. Bouin-fixed, tissue-mat embedded sections are stained in Pinkus' acid orcein for 1 1/2 hours and rinsed in distilled water. The sections are differentiated in 50% alcohol containing 1% hydrochloric acid, washed in tap and then in distilled water. The sections are next transferred for I to 2 minutes to the anilin blue, orange G, phosphomolybdic acid combination known as solution No. 2 of Mallory's connective tissue stain, diluted 1:1 with distilled water. They are then rinsed in distilled water, quickly passed into 95% alcohol, and dehydrated in absolute alcohol containing some orange G, after which they are cleared and mounted. Within less than two hours sections may be stained and mounted with the following results: elastic fibers — red; collagenic fibers — blue; muscle fibers — yellow; keratin — orange.     

The Use of Bismarck Brown Y in some new Stain Schedules

The staining quality of Bismarck brown Y may be improved and sterility maintained by adding 5% phenol to a 1% aqueous solution. Use the phenolic Bismarck brown in combination with iron alum hematoxylin except for stripped epidermis in the following procedures:

Stem and Root Schedule: Mordant sections from water in 4% iron alum for 10 minutes. Rinse in distilled water and stain in 0.5% aqueous hematoxylin for 1 minute or until darkly stained. Rinse in distilled water and destain in 2% iron alum until a gray color appears. Rinse thoroly in distilled water and intensify hematoxylin by transferring sections to 0.5% aqueous lithium carbonate until the desired black color appears. Rinse thoroly in distilled water and stain for 1-5 minutes in phenolic Bismarck brown. Rinse in distilled water, dehydrate successively in 30, 50, 70, 95 and 100% alcohol. Clear in methyl salicylate for 5 minutes, then to xylene for 3-5 minutes, and mount in balsam.

Middle Lamellae in Wood: Destain more thoroly in 2% iron alum than for the general stem and root schedule, and intensify in lithium carbonate for a longer period (about 1 hour).

White Potato Tuber Sections: Modify above schedule by reducing time of destaining in 2% iron alum to about 30-60 seconds and intensify hematoxylin until starch grains appear bluish in color. Stain in phenolic Bismarck brown for 1-2 minutes.

Wheat Grain Sections: Fix grain for sectioning when in “dough” stage. Use schedule the same as for potato tuber except for reducing time of staining in phenolic Bismarck brown to about 45 seconds.

Tradescantia zebrina Epidermis: Strip epidermis from leaf while submerged in water. Fix in 100% alcohol 10 minutes, pass thru 95, 70, 50, 30, and 10% alcohol to water. Stain in phenolic Bismarck brown for 10-20 minutes. Dehydrate, clear in methyl salicylate and mount in balsam.     

Improved nuclear staining with mordant blue 3 as a hematoxylin substitute.

For progressive staining 1 g mordant blue 3, 0.5 g iron a alum and 10 ml hydrochloric acid are combined to make 1 liter with distlled water. Paraffin sections are stained 5 minutes blued in 0.5% sodium acetate for 30 seconds and counterstained with eosin. For regressive staining, 1 g dye, 9 g iron alum and 50 ml acetic acid are combined to make 1 liter with distilled water. Staining time is 5 minutes followed by differentiation in 1% acid alcohol and blueing in 0.5% sodium acetate. Counterstain with eosin. In both cases results very closely results very resemble a good hematoxylin and eosin.     

Improved Nuclear Staining with Mordant Blue 3 as A Hematoxylin Substitute

For progressive staining 1 g mordant blue 3, 0.5 g iron alum and 10 ml hydrochloric acid are combined to make 1 liter with distilled water. Paraffin sections are stained 5 minutes, blued in 03% sodium acetate for 30 seconds and counterstained with eosin. For regressive staining, 1 g dye, 9 g iron alum and 50 ml acetic acid are combined to make 1 liter with distilled water. Staining time is 5 minutes followed by differentiation in 1% acid alcohol and blueing in 0.5% sodium acetate. Counterstain with eosin. In both cases results very closely resemble a good hematoxylin and eosin.     

Cleared Whole Mounts of Shoot Apices of Angiosperms for Topographic Study

Cleared and stained whole mounts of stem apices of two Labiates and of Phaseolus plumule giving a three-dimensional picture of the apical structure have been prepared as follows. Fix the buds in formalin-acetic acid-50% alcohol (5:5:90) for 24 hr or longer and then dissect under a binocular microscope to leave only the youngest leaves surrounding the apex. Wash for several minutes in distilled water and then clear the material in a 5% solution of sodium hydroxide at approximately 40° C for 24-48 hr. Wash thoroughly in several changes of distilled water, transfer to a solution of 1% tannic acid and 0.5% sodium salicylate for up to a minute. Wash briefly in distilled water and stain in a 1.5% solution of ferric chloride until blue-black. Wash in distilled water and dehydrate through 50%, 70%, 85%, 95% and 2 changes of absolute ethyl alcohol. If the xylem is not stained well, counter-stain for a few seconds in a 0.5% solution of safranin O in a 1:1 mixture of xylene and absolute alcohol and wash out the excess stain in the same mixture. Clear in 2 changes of xylene and place on a glass slide in thick Canada balsam. Orient with needles under low magnification and cover.     

The Staining of Radulae

A review of the various methods of staining and mounting radulae is given. Normally the radula should be extracted with 0.5 to 1% sodium hydroxide solution, and the associated tissues removed before staining. Two staining methods are recommended for facilitating the interpretation of radulae. Newly formed teeth and the bases of older ones are well stained by saturated aqueous chlorazol black E (up to 10 minutes). A more uniformly stained specimen) in which the cusps of all but the young teeth are alone stained, may be obtained by using the “oxidation-dahlia technic”. The radula is oxidized in N/10 potassium permanganate solution until black and subsequently decolorized in saturated aqueous oxalic acid. It is then stained in 0.1% aqueous dahlia (Hofmann's violet), the staining time varying from 10 to 30 minutes, according to the material. It is then dehydrated and passed through xylene and clove oil into Canada balsam. Other mountants may be employed but glycerin jelly is only recommended for the rapid examination of unstained radulae. Several other staining methods are mentioned, and general precautions to be observed while mounting are discussed.     

Destaining Agents for Iron Alum Hematoxylin

Detailed directions for using various destaining agents after iron alum hematoxylin are given. A saturated aqueous solution of picric acid is recommended for bringing out nuclei in whole mounts of protozoa. Differentiation with a mixture of 1 part 30% hydrogen peroxide, and 2 parts, 95% alcohol, is useful for cytoplasmic structures. Greater reliability and precision are claimed for these methods as compared with the conventional differentiation in iron alum.     

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.     

Aceto-Iron-Haematoxylin-Chloral Hydrate for Chromosome Staining

Aceto-iron-haematoxylin can be used combined with the clearing agent chloral hydrate for the squash method. The stain is prepared by dissolving 2 gm of chloral hydrate in 5 ml of a stock solution of 4% haematoxylin and 1% iron alum in 45% acetic acid, which has been allowed to ripen for 24 hr to 1 wk. Heat must not be used to hasten solution. The material (fixed in 1:3 acetic-alcohol) is put on a slide, the fixative removed and a drop of stain added; if necessary the material is crushed before the cover slip is placed in position. The preparations are now carefully heated until a slight colour change occurs. Squashing needs more pressure than in other techniques. This stain gives best results in zoological and botanical material not requiring hydrolysis, e.g., leucocytes, ascites cells, and cells undergoing spermatogenesis and microsporogenesis. Well-spread and selectively stained mitotic and meiotic figures can be obtained.     

Safranin and Anilin Blue with Delafield's Hematoxylin for Staining Cell Walls in Shoot Apexes

The following technic is suggested for staining cell walls in shoot apexes: After the usual preliminary steps through 50% ethyl alcohol, stain in 1 % safranin 0 for 24 hours. Rinse in tap water and place in 2% aqueous tannic acid for 2 minutes. After rinsing in tap water, stain for 2 minutes in 1 part Delafield's hematoxylin to 2 parts distilled water and rinse in tap water. Remove excess hematoxylin with acidified water (1 drop cone. HC1 in 200 ml. water), then place slides in 0.5% lithium carbonate for 5 minutes. Dehydrate through an ethyl alcohol series, then transfer from absolute alcohol to a saturated solution of anilin blue in “methyl cellosolve” for 5-10 minutes. Wash in absolute alcohol, rinse in a solution of 25% methyl salicylate, 33% xylene, 42% absolute ethyl alcohol and clear for 10 minutes in a solution of 2 parts methyl salicylate, 1 part xylene, 1 part absolute ethyl alcohol. Transfer through two changes of xylene and mount in “clarite” or suitable alternate. The resulting preparations will have clearly defined, dark-staining cell walls and will photograph well when “Super Panchro-Press, Type B” film (Eastman Kodak Co.) is used in conjunction with suitable Wratten filters.     

Chlorazol Black E as a Stain for Tension Wood

Sections containing gelatinous fibers were cut at 15 μ from material both fixed and stored in formalin-acetic-alcohol, 5:5:90 (of 70%). These sections were stained 5 min in a 1% aqueous solution of lignin pink (G. T. Gurr), differentiated quickly in water, soaked 5 min in 95% ethyl alcohol, dehydrated in absolute ethyl alcohol and counter stained 5 min with a 1% solution of chlorazol black E (G. T. Gurr) in methyl cellosolve, followed by dehydration in absolute ethyl alcohol, clearing in xylene and mounting in Canada balsam. The gelatinous layer was sharply defined as a dense black zone whilst the remainder of the cell wall stained light pink. The specificity of the technique was superior to that of safranin and light green, and was not easily obscured by overstaining. The technique is particularly useful for locating small zones of gelatinous fibres, and for photomicrographical work.     

Staining Nuclei and Chromosomes in Protozoa

Technics for free-living forms such as Paramecium and for parasitic forms such as the opalinid ciliates are described.

Paramecium: Fix paramecia in hot Schaudinn's fluid containing 5% of glacial acetic acid for 5-15 minutes. (A hot water bath for maintaining the proper temperature of the fixative is described.) Dehydrate up to 83% alcohol. Mount the specimens on albuminized cover glasses. (A table for mounting animals on cover glasses is described.) Apply a thin layer of collodion to the cover glass to prevent the loss of the specimens during the subsequent handling. Pass through descending grades of alcohol to water. Mordant in 4% iron alum for 24 hours. Stain in 0.5% hematoxylin for 24 hours. Destain in saturated aqueous picric acid. Rinse in tap water, expose to ammonia vapor for a second, and then rinse again in tap water. Wash in running water for 1 hour. Dehydrate. Clear, then mount in damar.

Opalinid Ciliates: Make smears on cover glasses and fix them while wet. If the opalinids are to be subsequently stained in hematoxylin, fix in hot Schaudinn's fluid (containing 5% of glacial acetic acid) for 5-15 minutes. Pass through descending grades of alcohol to water. Mordant in iron alum for 24 hours. Stain in hematoxylin for 24 hours. Destain in saturated aqueous picric acid. For Feulgen reaction, fix in a modified weak Flemming's fluid for 1 hour. Wash in running water for 30 minutes. Hydrolyze. Leave 3 hours in fuchsin decolorized with H₂SO₃ (Feulgen formula). Wash in H₂SO₃, then in running water for 15 minutes. Dehydrate up to 95% alcohol. Counterstain with fast green FCF for 2 minutes. Dehydrate in absolute alcohol. Clear, then mount in damar.