The Staining of Brunner's Gland and Other Neutral Mucins by Carmine, Hematoxylin and Orcein in Alkaline Solutions

Brunner's glands and other neutral mucins may be stained red, brownish red, and violet, respectively, by carmine, hematoxylin, and orcein from appropriate alkaline solutions. Carmine and hematoxylin in concentrations of 0.2-1% are dissolved in 60-70% alcohol containing 1% potassium carbonate; orein is used in a 0.2% alcoholic solution of sodium hydroxide. Staining times are 15 to 30 minutes. The stained sections are rinsed in 95% or absolute alcohol prior to xylene and mounting. The staining of these mucins is blocked by mild bromine oxidation. By using alcian blue 0.1% in 3% acetic acid for 5 minutes prior to the above stains, mucins may be characterized in the same preparation as acid, neutral or mixed.     

The staining of Brunner's gland and other neutral mucins by carmine, hematoxylin and orcein in alkaline solutions.

Brunner's glands and other neutral mucins may be stained red, brownish red, and violet, respectively, by carmine, hematoxylin, and orcein from appropriate alkaline solutions. Carmine and hematoxylin in concentrations of 0.2-1% are dissolved in 60-70% alcohol containing 1% potassium carbonate; orcein is used in a 0.2% alcoholic solution of sodium hydroxide. Staining times are 15 to 30 minutes. The stained sections are rinsed in 95% or absolute alcohol prior to xylene and mounting. The staining of these mucins is blocked by mild bromine oxidation. By using alcian blue 0.1% in 3% acetic acid for 5 minutes prior to the above stains, mucins may be characterized in the same preparation as acid, neutral or mixed.     

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 Selective Staining of Red Blood Cells

A method for the selective staining of red blood cells is described. Material is fixed in 10% neutral formalin in .85% NaCl and imbedded in paraffin or celloidin. Sections 6-10 μ are stained 1-5 minutes in chromotrope 2R. Basophilic and the less strongly acidophilic elements are decolorized with 5% phosphotungstic acid in 95% ethyl alcohol. Red blood cells and other strongly acidophilic elements that may be present in the preparation retain the chromotrope 2R. A counterstain of methyl blue may be used for staining the decolorized basophilic elements. As a result, erythrocytes are stained red by the chromotrope 2R, and basophilic elements blue, by the methyl blue. Less strongly acidophilic elements, having little affinity for either primary or secondary dye, are colorless or gray.     

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.     

New Fuchsin-Hematoxylin-Eosin; A Stain for Acid-Fast Bacilli and Surrounding Tissue

This is a staining technique for histopathologic evaluation of tissue reaction in the environs of acid-fast tubercle bacilli (avian and bovine) in sections. Fresh tissue is fixed in 10% neutral formalin and processed in the usual manner for embedding in paraffin. Sections are cut approximately 6 μ. thick, dewaxed, hydrated, and stained with Harris' hematoxylin. They are rinsed in tap water, differentiated in add alcohol, washed in tap water, given a distilled water rinse and stained at 20-30° C in a 1% solution of new fuchsin in 5% phenol. Each slide is then handled individually by placing it directly into a saturated aqueous solution of Li₂CO₃ and agitated gently for a few seconds. This is followed by differentiation with 5% glacial acetic acid in absolute or 95% ethyl alcohol until the color stops running. Two rinses in absolute or 95% ethyl alcohol follow. The sections are then counterstained in the color add of eosin Y prepared according to the method of Schleicher (Stain Techn., 28, 119-23, 1953) and used as an 0.025% solution in absolute alcohol. Following passage through 2 changes of absolute alcohol, the sections are cleared in xylene, then mounted in Permount or similar synthetic resin. The add-fast barilli are emphasized by their bright retractile red color within a contrasting background of hematoxylin and eosin.     

New Fuchsin-Hematoxylin-Eosin; A Stain for Acid-Fast Bacilli and Surrounding Tissue

This is a staining technique for histopathologic evaluation of tissue reaction in the environs of acid-fast tubercle bacilli (avian and bovine) in sections. Fresh tissue is fixed in 10% neutral formalin and processed in the usual manner for embedding in paraffin. Sections are cut approximately 6 μ. thick, dewaxed, hydrated, and stained with Harris' hematoxylin. They are rinsed in tap water, differentiated in add alcohol, washed in tap water, given a distilled water rinse and stained at 20-30° C in a 1% solution of new fuchsin in 5% phenol. Each slide is then handled individually by placing it directly into a saturated aqueous solution of Li₂CO₃ and agitated gently for a few seconds. This is followed by differentiation with 5% glacial acetic acid in absolute or 95% ethyl alcohol until the color stops running. Two rinses in absolute or 95% ethyl alcohol follow. The sections are then counterstained in the color add of eosin Y prepared according to the method of Schleicher (Stain Techn., 28, 119-23, 1953) and used as an 0.025% solution in absolute alcohol. Following passage through 2 changes of absolute alcohol, the sections are cleared in xylene, then mounted in Permount or similar synthetic resin. The add-fast barilli are emphasized by their bright retractile red color within a contrasting background of hematoxylin and eosin.     

Evaluation of improved zirconyl hematoxylin compared to the classical pH 2.5 Alcian blue method for demonstrating acid mucins

Abstract

Acid mucins have diagnostic significance for many pathological conditions, especially in certain tumors. We compared the classical pH 2.5 Alcian blue method to a new, improved zirconyl hematoxylin (IZH) method for demonstrating acid mucins using two fixatives: Bouin`s solution and 10% neutral buffered formalin (NBF). We used rabbit small intestine, large intestine and trachea. Specimens were fixed in Bouin`s solution and NBF. A total of 160 paraffin sections were prepared and stained with pH 2.5 Alcian blue and IZH. The stained acid mucins were assessed using digital image analysis software. Stained mucins were quantified for each staining procedure and fixative. No important differences were observed in acid mucin staining by either method after either fixative. The IZH method provides results as good as pH 2.5 Alcian blue and can be used to obtain reliable staining for acid mucins.     

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.     

Evaluation of improved zirconyl hematoxylin compared to the classical pH 2.5 Alcian blue method for demonstrating acid mucins

Acid mucins have diagnostic significance for many pathological conditions, especially in certain tumors. We compared the classical pH 2.5 Alcian blue method to a new, improved zirconyl hematoxylin (IZH) method for demonstrating acid mucins using two fixatives: Bouin`s solution and 10% neutral buffered formalin (NBF). We used rabbit small intestine, large intestine and trachea. Specimens were fixed in Bouin`s solution and NBF. A total of 160 paraffin sections were prepared and stained with pH 2.5 Alcian blue and IZH. The stained acid mucins were assessed using digital image analysis software. Stained mucins were quantified for each staining procedure and fixative. No important differences were observed in acid mucin staining by either method after either fixative. The IZH method provides results as good as pH 2.5 Alcian blue and can be used to obtain reliable staining for acid mucins.     

Quad-Type Stain for the Simultaneous Demonstration of Intracellular and Extracellular Tissue Components

This technic for the simultaneous demonstration of several different tissue components works equally well on invertebrate and vertebrate tissue if they have been treated with nonchromate fixatives Sections 4-7 μ thick are stained 30 min in 1% Alcian blue, then treated with alkaline alcohol for 2 hr. They are stained in Verhoeff's hematoxylin for 4-6 hr, and rinsed in alcohol; stained in woodstain scarlet-acid fuchsin for 3 min, decolorized in 5% phosphotungstic acid for 20 min and finally stained 5-8 min in alcoholic saffron. Collagen and bone are stained yellow; elastin, myelin and nucleic acids, purple to black; muscle, chitin, cytoplasm, fibrinoid and acid secretion, bright red to lavender; ground substances and mucus, blue-green. Fibrous connective tissue, cartilage, bone and glandular epithelia are exceptionally well demonstrated by this method. Slides stained in this manner are well suited for color photomicrography and as demonstrations in the classroom.     

Stain Technic for Nissl's Granules Following Alcohol-Formol Fixation

1. Nerve tissue is fixed 2-4 hrs in a 5% solution of strong formalin in commerical 95% alcohol.

2. If dehydration is perfect, either chloroform or xylol may be used as a clearing agent.

3. A slow method of paraffin infiltration is advisable.

4. Sections should be cut 10-12 microns in thickness.

5. Coplin staining jars should be annealed by placing them on a rack in a pan of cold water, bringing the water to the boiling point, and allowing the jars to stand in boiling water for twenty minutes.

6. One per cent aqueous solutions of either methylene blue or Grübler's Neutral Roth are used as specific stains for Nissl's granules.

7. These stains are heated to boiling in a beaker, the slides are placed in the Coplin jars which are partially submerged in boiling water, and the hot stain poured into the jars. The flame beneath the water bath is turned down and the slides left for 20 minutes.

8. The excess of primary stain is washed off in 25% and 50% alcohol and the slides passed rapidly thru the alcohol series to absolute alcohol, and finally to xylol.

9. When counterstaining is desired, nigrosin in 1% aqueous solution, methyl orange, saturated solution in 50% alcohol, or, a 0.5% solution of eosin in 50% alcohol are recommended. These stains are used cold, and the slides are merely dipped in them after the excess of primary stain has been washed out in 25% and 50% alcohol.

10. If a cold primary stain is desired, a saturated solution of thionin in distilled water, acidified with 1% carbolic acid, will prove specific for the Nissl substance. Sections should be stained 5–10 minutes in thionin, then passed rapidly thru to absolute alcohol, and xylol. The same counterstains may be used as in the hot method.

11. Sections prepared by the hot method show little tendency to fade after ten years use.

12. Excepting neutral red, all the stains used in this technic are carried by the National Aniline and Chemical Company and are satisfactory. Coleman and Bell neutral red may be substituted for Grübler's Neutral Roth with good results.     

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.     

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 Use of Buffered Solutions in Staining: Theory and Practice

The importance of pH in staining tissue is emphasized. The effect of pH upon the selectivity and intensity of staining with iron hematoxylin, malachite green, and eosin Y is considered. Many difficulties may be avoided by staining in the higher alcohols and directions are given for the preparation of buffer solutions from pH 1.2-8 in alcohol. The concentration of stains, time of staining, and order of staining are discussed for progressive and regressive staining. At pH 8 in 95% alcohol very few tissues stain with malachite green at a concentration of 1/1000 saturated. At pH 6 most cytoplasmic elements stain with malachite green at a concentration of 1/1000 saturated or with eosin Y at 1/250 saturated. As the pH is lowered more tissue elements stain until the nucleus is completely stained. This behavior is in accord with the theory of chemical combination of dyes with proteins, which states that proteins combine with basic dyes on the basic side of their isoelectric points and with acid dyes on the acid side of their isoelectric points. With hematoxylin stain the pH range is much shorter. A satisfactory hematoxylin stain is composed of 0.1% hematoxylin, 0.1% FeCl₃, and HCl to bring the pH to 1.2-1.6 in 80% alcohol. With this stain, which may be used immediately, the nuclei of most tissues begin to stain at pH 1.2 and much of the cytoplasm will be stained if the pH is raised to 1.4. The shortness of this effective pH range is thought to be due to the dissociation of the hematoxylin-iron-protein complex. The use of different dyes successively at different pH values, such as hematoxylin at 1.3, malachite green at 8, and eosin at 6, permits better differentiation of the tissue elements, and intelligent variations in the staining technic.     

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.     

Block staining of mammalian tissues with hematoxylin and eosin

Various mammalian tissues were stained en bloc with hematoxylin and eosin after fixation and prior to embedding in paraffin wax and sectioning. The choice of fixative is important and best results are obtained using Worcester's Fluid, a combination of saturated aqueous mercuric chloride, formaldehyde, and glacial acetic acid. After fixation, blocks of tissue up to 1.5 cm thick are stained for seven days in hematoxylin. Excess stain is removed by washing tissues in running water overnight. Tissue blocks then are dehydrated with graded concentrations of ethyl alcohols to 80% and counterstained, with further dehydration, in 0.5% spirit soluble eosin in 90% ethyl alcohol for five days. The tissue is subsequently transferred to 90% ethyl alcohol overnight to differentiate eosin staining; dehydration is completed in absolute ethyl alcohol. The blocks are cleared in in cedarwood oil and briefly in xylene prior to embedding, sectioning, and mounting. Following removal of wax by xylene, coverslips are applied. General morphological and histological features were particularly well differentiated and very selectively and reliably stained by this method.     

Block Staining of Mammalian Tissues with Hematoxylin and Eosin

Various mammalian tissues were stained en bloc with hematoxylin and eosin after fixation and prior to embedding in paraffin wax and sectioning. The choice of fixative is important and best results are obtained using Worcester's Fluid, a combination of saturated aqueous mercuric chloride, formaldehyde, and glacial acetic acid. After fixation, blocks of tissue up to 1.5 cm thick are stained for seven days in hematoxylin. Excess stain is removed by washing tissues in running water overnight. Tissue blocks then are dehydrated with graded concentrations of ethyl alcohols to 80% and counterstained, with further dehydration, in 0.5% spirit soluble eosin in 90% ethyl alcohol for five days. The tissue is subsequently transferred to 90% ethyl alcohol overnight to differentiate eosin staining; dehydration is completed in absolute ethyl alcohol. The blocks are cleared in cedarwood oil and briefly in xylene prior to embedding, sectioning, and mounting. Following removal of wax by xylene, coverslips are applied.

General morphological and histological features were particularly well differentiated and very selectively and reliably stained by this method.     

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.     

Delafield'S Hematoxylin and Safranin for Staining Plant Materials

An improved schedule is suggested for staining plant materials in Delafield's hematoxylin and safranin. Tissues are stained first in Delafield's hematoxylin. A short bath in acidulated water (1 or 2 drops concentrated HCl to 100 cc.) removes objectionable precipitates, and at the same time serves as a destaining agent. The acid bath must be followed quickly by a thoro wash in tap water, or dilute lithium carbonate solution, to restore the original dark blue color (made reddish in the acid bath) of the hematoxylin and to “set” the stain. Once the hematoxylin solution is satisfactory, none of the reagents ordinarily used will remove it—unless they contain acid. Tissues are counterstained in rapid safranin (5 drops analin in 100 cc. of 1% safranin 0 in 50% ethyl alcohol); this materially lessens the time necessary for staining. The safranin is de-stained in 50% ethyl alcohol (which does not affect the hematoxylin) until sharp differentiation is secured. If destaining is too slow, or differentiation poor, a quick rinse in acidulated 50% alcohol usually sharpens contrast of the stains. This must be followed quickly by a wash in 50% alcohol containing lithium carbonate to neutralize the acid. Dehydrate, and mount as usual. This schedule allows each stain to be individually, and independently, controlled at the will of the operator.