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.     

The Combined Perls-Hematoxylin-Eosin Stain

To provide a routine check for the presence of ferric iron in sections, Perls' method was combined with hematoxylin and eosin as follows. Deparaffinized sections of formalin-fixed tissues are stained in Perls' reagent (1:1 2%, w/v, of potassium ferrocyanide in distilled water and 2%, v/v, concentrated HCl in distilled water) for 20 min. After brief rinsing in distilled water stain sections in Mayer's hemalum, wash in tap water for 5 min, counterstain in 0.5% (w/v) eosin B in 50% ethyl alcohol for 15 sec. Rinse in tap water, dehydrate and mount as usual.     

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.     

Hematein-its Advantages for General Laboratory Usage

The advantages of hematein over hematoxylin are that it is easy to prepare, easy to use, and saves time; while it gives equally good results. The writer has been employing for some time a pre-war imported hematein and until within the last few months has been unable to locate a satisfactory product of recent manufacture, either domestic or foreign. At the request of the Biological Stain Commission, however, an American manufacturer has at last put on the market a C. P. hematein which gives splendid results. The technic is as follows:

Paraffin or celloidin sections of Bouin or Zenker-formol material are run down to water and stained about 5 minutes in Mayer's hemalum (0.5 g. hematein ground up in a glass mortar with 10 cc. 95% alcohol and added to 500 cc. of 5% aqu. sol. potassium alum.) Rinse 1 to 3 seconds in tap water. Dip 1 to 3 seconds in eosin B (1 part 0.5% sol. in 20% alc. added to 2 pats dist. water; filtered from time to time). Wash several minutes in running water or in several changes of tap water. Dehydrate and mount; with unattached celloidin sections this may be done by running up to 95% alcohol, spreading on slide, blotting, wetting with absolute alcohol, draining and mounting in euparal.     

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.     

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.     

A Staining Combination for Phloem and Contiguous Tissues

A technic is described for producing critically stained preparations of phloem tissue. The preparations promise to be relatively stable. Sections of fixed unembedded or of embedded (paraffin or celloidin) phloem, cambium, and xylem are (1) stained in Foster's tannic acid-ferric chloride combination; (2) treated with 1% NaHCOg in 25% or 50% ethyl alcohol for 30 minutes; (3) stained in a saturated solution of lacmoid (made alkaline by adding a few ml. of 1% NaHCO₃ in 25% alcohol) for 12 to 18 hours; (4) dehydrated and cleared in a series composed of 1% solution of NaHCO_s in 50% ethyl alcohol, 80%, 95%, and absolute alcohol, equal proportions of absolute alcohol, clove oil, and xylene, and finally pure xylene; and (5) mounted in a neutral resin. Callose and lignified secondary walls are blue or blue-green in color, cellulose walls and stainable protoplasmic contents are generally light brown. The technic has been successful with sections from 5 to 40μ in thickness, and the staining has been satisfactory for both color and black and white photomicrography.     

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.     

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.     

Application of the Papanicolaou Stain to Paraffin Sections

Routine paraffin sections from tissues fixed either in aqueous formalin, 80% alcohol (with or without 1% trichloracetic acid added), Carnoy's alcohol-chloroform-acetic (6:3:1) and Bouin's fixative were stained as follows: Harris' hematoxylin, 6 min; running water, 2-3 min; ascending grades of alcohol to 95%; orange G, 0.5% and phosphotungstic acid, 0.015% in 95% alcohol, 5 min; 95% alcohol, 2 changes; Papanicolaou's EA36, 2.5 min; dehydration, clearing, and covering in Permount. The results show morphology better than hematoxylin and eosin and the technic is recommended particularly for keratin, which always stains bright orange.     

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.     

A consistent phosphotungstic acid hematoxylin stain for glial fibers.

After deceration, celloidinization and hydration, oxidize 10 micron paraffin sections for 15 min in a solution containing 0.3 g KMnO4 and 0.1 ml conc. H2SO4 per 100 ml distilled water. Wash in water and reduce in 5% oxalic acid until the sections are colorless. Wash thoroughly in water and place in 4% iron alum solution for two hours. Wash briefly in water and stain for two hours in phosphotungstic acid hematoxylin. Rinse briefly in 95% ethanol and dehydrate in n-butyl alcohol or absolute ethanol for 4 min with two changes, clear and mount. Glial fibers, myofibrils, red blood cells, etc. are stained blue while astrocyte cell bodies, collagen, etc. are stained red. This stain has proven highly consistent in a wide variety of astrocytic derangements. Despite the intensity of this PTAH modification, false positive staining was not observed.     

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.     

Staining Methods For Studying Ingredient Distribution In Baked Cakes

A method is described for preparing cake crumb for sectioning and staining. Previous to embedding, the fat was stained and fixed by exposing small blocks of cake to the fumes from a 5%, freshly-prepared, aqueous solution of osmic acid (OsO₄). This was followed by dehydration in ethyl alcohol and tertiary butyl alcohol, removal of air under vacuum and infiltration with paraffin.

Sections were cut 20 and 9Op thick and mounted with water.

Wax was removed by immersion in xylene. The sections were rehydrated in a series of ethyl alcohol dilutions, from concentrated to dilute, then transferred to distilled water.

Protein was then stained pink by immersion of the slides in an acidified 0.04% water solution of eosin Y, or starch was stained blue with a dilute aqueous solution of iodine. Ten grams iodine and 10 g. KI were dissolved in 25 ml. distilled water. This stock solution was diluted for use one to two hundred times.

The relationship between protein and starch was demonstrated by staining the sections with eosin, differentiating in 50% alcohol and staining with iodine.

When slides of cake crumb were prepared in this way, the fat was stained black, the protein bright pink and the starch granules a dark blue.     

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.     

Effect of phosphomolybdic acid on the binding of Sudan black B

Paraffin sections of tissues fixed in absolute alcohol or Carnoy's fluid were mordanted in a 1% aqueous solution of phosphomolybdic acid, stained in saturated solutions of Sudan black B, acetylated Sudan black, various solvent and basic dyes in 70% ethyl alcohol for 5 min at room temperature, dehydrated in alcohol and covered in Permount. Sudan black B and other dyes with basic groups stained basement membranes, reticulum and collagen fibers intensely. Acetylated Sudan black, Sudan IV and oil red 0 did not color any tissue structures. Control sections, without pretreatment, did not bind Sudan black B. These findings indicate interaction between basic groups of the dye and free acid groups of phosphomolybdic acid.     

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.     

A Consistent Phosphotungstic Acid Hematoxylin Stain for Glial Fibers

After deceration, celloidinization and hydration, oxidize 10 micron paraffin sections for 15 min in a solution containing 0.3 g KMnO₄, and 0.1 ml conc. H₂SO₂, per 100 ml distilled water. Wash in water and reduce in 5% oxalic acid until the sections are colorless. Wash thoroughly in water and place in 4% iron alum solution for two hours. Wash briefly in water and stain for two hours in phosphotungstic acid hematoxylin. Rinse briefly in 95% ethanol and dehydrate in n-butyl alcohol or absolute ethanol for 4 min with two changes, clear and mount. Glial fibers, myofibrils, red blood cells, etc. are stained blue while astrocyte cell bodies, collagen, etc. are stained red. This stain has proven highly consistent in a wide variety of astrocytic derangements. Despite the intensity of this PTAH modification, false positive staining was not observed.     

A Nissl Method Using Buffered Solutions of Thionin

Experiments were performed in an attempt to obtain a rapid method for staining the chromophilic substance of formalin-fixed nerve cells differentially without resorting to over-staining and subsequent acid-decolorizing. A satisfactory procedure using thionin in dilute buffered solutions was developed: Prepare a stock solution of the dye using 1 g. in 100 ml. of distilled water. Prepare veronal-acetate buffers (Michaelis, 1931) in the range of pH 5 to pH 3.S. To each 10 ml. of the buffer add 0.5 ml. of the stock dye solution. After rinsing in CO₂-free distilled water place mounted or unmounted sections in this solution. (Freshly fixed material, 10μ to 20μ thick, is completely stained in 10 to 20 minutes but over-staining does not occur when longer times are allowed.) Return sections to distilled water (2 changes) and wash until diffusion of excess dye is no longer visible. Wash in 70% ethyl alcohol (2 changes) until diffusion of excess dye is no longer visible. Dehydrate in 95% ethyl alcohol and normal butyl alcohol, clear and mount.

Optimum staining of chromophilic material occurs at pH 3.65. Glial processes are well stained at pH 4.6. Nissl bodies and glial cytoplasm are purplish blue, nuclear chromatin is blue, background is clear at pH 3.65 but pale blue at pH 4.9.