Delafield's Hematoxylin and Safranin for Staining Meristematic Tissues

The following technic is suggested for staining permanent preparations of meristematic tissues: Prepare and mount the sections by the usual paraffin method. From water, stain them 2-10 minutes in a solution made by adding 2-4 cc. of Delafield's hematoxylin to a Coplin jar full of tap water. As staining is progressive, the sections should be examined from time to time with a microscope. When the cell walls have become a deep purple, transfer the preparations, thru the usual series, to a mixture of xylol-absolute-alcohol in equal parts, and from this to a counterstain made by adding 4-6 cc. of a saturated solution of safranin in absolute alcohol to a Coplin jar full of xylol (75%) with absolute alcohol (25%). This stains the nuclei. Leave the sections in the counterstain at least 2 hours and then rinse them in xylol-absolute-alcohol (1:1) to remove excess safranin. Transfer them to pure xylol and then mount them in neutral balsam.     

Frozen Section Micro-Incineration

A method for micro-incineration of frozen sections is described. Material containing diffusible or soluble salts is cut on the freezing microtome and the sections are placed into xylol and mounted out of xylol onto Corex D slides previously filmed with glycerin-gelatin medium. Material containing non-diffusible or insoluble salts can be fixed in 10% formalin before sectioning. Sections of the fixed material are dehydrated thru 50, 70, and 95% ethyl alcohol and mounted out of absolute alcohol onto Corex D slides previously fumed with glycerin-gelatin medium. After mounting by either procedure the sections are incinerated in an electric furnace and the temperature of incineration is dependent on the type of tissues to be incinerated and the character of the salts present. The method is time saving and when no fixation is required the whole procedure can be carried out in one hour.     

New Rapid Methods for Tissue Diagnosis

Two new methods applicable to the staining of fixed and fresh frozen tissue sections are presented herein. In addition certain improvements are suggested for the technic reported by Geschickter, Walker, Hjort and Moulton (1931). In brief the procedures are as follows:

The thionin eosinate method of Geschickter et al (1931). This procedure has been modified as follows:

A mixture of diethylene glycol, 40 parts, ethylene glycol, 40 parts, and grain alcohol, 20 parts is superior to ethylene glycol, 80 parts, and ethyl alcohol, 20 parts, as a solvent for the compound stain in that the staining is intensified.

Ethylene glycol monobutyl ether supplants diethylene glycol monobutyl ether because of its lower viscosity.

Ethyl phthalate replaces butyl phthalate on account of a more satisfactory viscosity.

The methyl green eosinate procedure is the same as the modified thionin eosinate method except for the following variations:

The staining time is increased to one minute.

Decolorization and washing are reduced to about 15 seconds.

The hematoxylin-eosin method. After cutting, the tissue sections are carried thru the following steps:

Unfold in water; transfer to formalin (4 to 40%) for at least 30 seconds; stain in hematoxylin (Harris) for 30 to 60 seconds; wash in water, 5 seconds; decolorize in 0.1% HC1 or saturated aqueous picric acid, 5 seconds; wash in water, S seconds; float in 0.5% ammonia, 5 to 10 seconds; wash in water, 5 seconds; stain in 5% aqueous eosin, 15 seconds¹; wash in water, 5 to 10 seconds; dehydrate in a mixture of diethylene glycol, 30 parts, and ethyl alcohol, 70 parts, 5 to 10 seconds; dehydrate in ethylene glycol monobutyl ether, 5 to 10 seconds; clear in ethyl phthalate, 5 to 10 seconds; float on a glass slide, blot with photographic lintless blotter, place a drop of neutral gum damar on the section, and cover with glass cover slip.     

A Method for Making Aceto-Carmin Smears Permanent

Anthers are collected and placed in a solution of 1 part acetic acid to 3 parts of absolute alcohol. The contents of the anther are squeezed out on a slide in a drop of Belling's iron-aceto-carmin solution and a cover glass placed over the drop. Care should be taken to remove all anther walls and flower parts. Heat the slide over an alcohol flame for a second, repeating 4 or 5 times. Place the slide in a petri dish filled with a 10% solution of acetic acid. When the cover glass has risen away from the slide gently remove the cover glass and place in a Coplin jar containing equal parts of alcohol and acetic acid. Likewise, place the slide in this solution. Run both cover and slide thru the following solutions: 1 part acetic acid to 3 parts absolute alcohol, 1 part acetic acid to 9 parts absolute alcohol, absolute alcohol and finally equal parts of absolute alcohol and xylol. Recombine the cover and slide in xylol-balsam directly from this solution.     

A Method for Making Aceto-Carmin Smears Permanent

Anthers are collected and placed in a solution of 1 part acetic acid to 3 parts of absolute alcohol. The contents of the anther are squeezed out on a slide in a drop of Belling's iron-aceto-carmin solution and a cover glass placed over the drop. Care should be taken to remove all anther walls and flower parts. Heat the slide over an alcohol flame for a second, repeating 4 or 5 times. Place the slide in a petri dish filled with a 10% solution of acetic acid. When the cover glass has risen away from the slide gently remove the cover glass and place in a Coplin jar containing equal parts of alcohol and acetic acid. Likewise, place the slide in this solution. Run both cover and slide thru the following solutions: 1 part acetic acid to 3 parts absolute alcohol, 1 part acetic acid to 9 parts absolute alcohol, absolute alcohol and finally equal parts of absolute alcohol and xylol. Recombine the cover and slide in xylol-balsam directly from this solution.     

The Smear Technic in Plant Cytology

The following method of making permanent smears of pollen mother cells is in general use and gives excellent results. Determine the stage of meiosis from aceto-carmin mounts. Smear the pollen mother cells on a dry slide. Fix in Navaschin's or a modified Flemming's solution from 1 to 2 hours. Wash in 10 to 20% alcohol from 15 to 30 minutes. Stain in 1% aqueous crystal violet from 1 to 5 minutes. Rinse in water and pass thru 30 to 50% alcohol, about 15 to 20 seconds in each. Transfer to 80% alcohol containing 1% iodine and 1% potassium iodide for 30 seconds. Destain with absolute alcohol, followed by clove oil. xylol, balsam and cover.

Permanent smears for chromosome counts can be quickly made by smearing pollen mother cells on a dry slide, fix and stain with aceto-carmin, dehydrate with mixtures of absolute alcohol and acetic acid, follow with xylol, balsam, and cover.     

A Tri-Basic-Dye Stain for Nerve Cells

A new staining method has been developed for the study of nerve cells and Nissl granules which combines three basic dyes, cresylecht violet, toluidine blue and thionin. The use of this tri-basic-dye stain results in finished preparations that are critically stained and permanent. Paraffin sections (4 μ sections preferably) are mounted on slides by the starch medium, deparaffinized and stained by the tribasic staining solution. After differentiation in acidified distilled water, sections are dehydrated, returned to stain solution and again dehydrated, then cleared and mounted in Clarite. Various vertebrate material including normal and pathological human tissues have been stained with this triple dye solution. Especially for pathological material, re-immersion of slides in the staining and 80% alcohol solutions before mounting, differentially intensifies the staining reaction. Fixatives used were 10% formalin, 95% alcohol, Bouin and formalin-Bouin (10% formalin followed by Bouin).     

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.     

Novel Fixation of Plant Tissue, Staining through Paraffin with Alcian Blue and Hematoxylin, and Improved Slide Preparation

Onion (Allium cepa) root tips were fixed in a proprietary solution without aldehyde, toxic metals or acetic acid. Fixed specimens were embedded in paraffin, sectioned on a rotary microtome and mounted on detergent-washed slides without adhesive. Slides with ribbon segments affixed were immersed in 0.2% aqueous alcian blue 8GX in screw-capped Coplin jars in a water bath at 50 C for 1 hr. Excess alcian blue was rinsed off under cold running tap water and the slides were immersed in quick-mixed hematoxylin at room temperature for 15 min. Stained slides were deparaffinized, rinsed with isopropanol, air dried, and coverslips were affixed with resin. Thus, the traditional paraffin microtechnique has been modified at all steps from fixation to finishing slides with coverslips.     

A Simplified Nissl Stain with Thionin

Recently two articles on the use of thionin as a cell stain for neurological materials have appeared. One utilizes a solution buffered in the acid range³; the other uses a “steaming” staining solution⁴. For some time we have been using thionin as a routine stain after either formalin or alcohol fixation and our method is so simple and has given such satisfactory results with a variety of brands of thionin that it seemed to be worthy of more general use. Briefly the method consists of placing the celloidin sections in a 0.05% solution of Li₂CO₃ (the percentage of Li₂CO₃ is non-critical) for about 5 minutes and then grossly overstaining in a 0.25% solution of thionin in a 0.05% solution of Li₂CO₃ in distilled water. The overstaining is necessary if all the stain is to be removed from the background. The sections are then passed through distilled water, 70 or 80% alcohol, two changes of butyl alcohol, two changes of xylene and mounted with Clarite. For most material, split mica cover-slips are quite satisfactory. The time of differentiation may be considerably lessened by the use of the differentiator recommended by Neumann (1942) except that we find the chloroform superfluous and transfer the sections to the aniline solution from 95% alcohol. Less fading seems to occur if the aniline differentiator is followed by a saturated solution of Li₂CO₃ in 95% alcohol.     

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 Portable Rack for Staining Jars

A simple and inexpensive rack has been designed for the transport and storage of stains and reagents in Coplin staining jars in bacteriological and histological laboratories. The rack promotes ease of carrying, prevents spillage, and keeps the jars permanently in the correct sequence. Details of construction are given. The design can be modified to hold as many jars as necessary and can also be modified to accommodate Stender or other type staining jars or reagent bottles.     

Stain Technology a Bone Stain for Osteoid Seams in Fresh, Unembedded, Mineralized Bone

Fresh, ground, mineralized bone sections 75-100 μ thick are stained 90 minutes or 48 hours in the Bone Stain, a preparation containing fast green FCF, orange G, basic fuchsin, and azure II. Surface stain is then removed by grinding under running water. Sections are washed in 0.1% zephiran chloride (benzalkonium chloride) or in 0.01% mild soap and again washed in tap water, followed with distilled water. Sections are next differentiated in 0.01% acetic acid in 95% methanol, dehydrated in 95% ethanol and 100% ethanol, cleared in alcohol:xylene 1:1, 1:4, 1:9 and 2 changes of xylol, and then mounted permanently in Eukitt's mounting media.

Osteoid seams stain either green to jade green or red to dark red, incompletely mineralized bone red or orange yellow, and the zone of demarcation light green. The walls of lacunae, canaliculae, feathered bone, procedural artifacts and periosteocyte lacunar low-density versions stain red.

The method helps in the differential diagnosis of certain metabolic bone diseases in human biopsy and autopsy material.     

Further Studies on a Modification of the Gram Stain

In perfecting the modification of the Gram-stain previously proposed, the following points are of interest:

1. Acetone is too strong a decolorizer for Gram-positive organisms and alcohol too weak for Gram-negative organisms. Consequently, it is now recommended that equal parts of acetone (100% c.p.) and ethyl alcohol (95%) be used as a decolorizing agent. The time of application should not ordinarily exceed 10 seconds.

2. Aqueous basic fuchsin (0.1%) serves as a strongly contrasting counterstain. Prolonged application renders Gram-positive organisms doubtful or Gram-negative, while short application renders Gram-negative organisms doubtful or Gram-positive. Twenty (20) seconds is therefore recommended as the time of application of the counterstain.

3. The method here described, with due regard for its limitations, is of value in Gram-staining pure or mixed cultures as well as for organic materials, such as Acidophilus milk, feces, etc., either for research purposes or classroom use. The method is as follows:

Air-dry film and fix with least amount of heat necessary.

Flood with dye for 5 minutes. Previously mix 30 drops of a 1% aqueous solution of crystal violet or methyl violet 6B with 8 drops of a 5% solution of sodium bicarbonate. Allow the mixture to remain for 5 minutes or more.

Flush with iodine solution for 2 minutes. Two grams iodine dissolved in 10 cc. normal sodium hydroxide solution and 90 cc. water added.

Drain without blotting but do not allow film to dry.

Add a mixture of equal parts of acetone and alcohol drop by drop until the drippings are colorless. (10 seconds or less.)

Air-dry slide.

Counterstain for 20 seconds with 0.1% aqueous solution of basic fuchsin.

Wash off excess stain by short exposure to tap water and air-dry. If slide is not clear immersion in xylol is recommended.     

Staining Procedures for Ultrathin Sections of Tissues Embedded in Polyester Resin

Tissues were fixed for 30 min In cold (0-2° C) 1% OsO₄ (Palade) buffered at pH 7.7, to which 0.1% MgCl₂ was added. Dehydration was in a graded ethanol series (containing 0.5% MgCl₂) at 0-2° C, and terminated with 2 changes of absolute ethanol. Tissues were then transferred by a graded series to anhydrous acetone. Infiltration of the tissue with Vestopal-W (a polyester resin), is gradual with the aid of graded solutions of Vestopal-W in acetone. The infiltrated tissue is encapsulated and initial polymerization is done under ultraviolet light at room temperature for 8-16 hr. This is followed by final hardening at 60° C for 36-48 hr. Sections (0.2-1 μ) were cut, dried on slides, placed in acetone for 1 min and then treated by either of the following staining procedures: (1) Thionin-azure-fuchsin staining: Flood the preparation with 0.2% aqueous thionin and heat to 60-80° C for 3 min; if the preparation begins to dry, add stain. Rinse in distilled water. Flood the slide with 0.2% azure B in phosphate buffer at pH 9. Heat to 60-80° C for 3 min; do not permit the preparation to dry. Rinse in distilled water. Dip the slide in MacCallum's variant of Goodpasture's carbol-fuchsin stain for 1-2 sec. Rinse in distilled water. Check the preparation microscopically for intensity of the fuchsin stain. Repeat dips as may be needed to obtain the desired intensity. Rinse in distilled water. Dehydrate quickly in 95% and absolute alcohol; clear in 2 changes of xylene and cover in Permount or similar synthetic resin. (2) Thionin-azure counterstain for the periodic acid-Schiff reaction: Oxidize the tissue in 0.5% periodic acid for 15 min and transfer to Schiff's leucofuchsin solution for 30 min. Counterstain with 0.5% aqueous thionin for 3 min; wash in distilled water; stain in 0.2% azure B in phosphate buffer at pH 5.5; wash in distilled water; dehydrate; clear and cover as in the first method. For temporary preparations let dry after absolute alcohol and apply a drop of immersion oil directly on the section.     

Differentiation of Two Classes of Acidophiles in the Anterior Pituitary of the Female Rabbit and Cat

A differential stain for the anterior pituitary of mammals, based directly on Heidenhain's 'azan' modification of Mallory's connective tissue stain has been devised. Tissue is fixed for 24 hours in a saturated solution of corrosive sublimate in physiological saline (90 parts) and formalin (10 parts) and washed directly in 70% alcohol for 48 hours. Sections are treated on the slide with a 3% solution of potassium bichromate for 12 hours. Two classes of acidophiles are demonstrated: one which stains selectively with azocarmine; and the ordinary acidophile which stains with orange G. The special acidophile has been demonstrated in the female rabbit and cat but has not been found in the mouse or rat.     

Basic Two-Dye Stains for Epoxy-Embedded 0.3-1 μ Sections

Dyes used in the 3 methods recommended are: I, thionin and acridine orange (T-AO); II, Janus green and Darrow red (JG-DR); III, methyl green and methyl violet (MG-MV). The first 2 methods were two-solution stains, applied in sequence; the third, required only one solution since methyl violet is present in commercial methyl green. Staining solution and timing was as follows: Method I. 0.1% thionin in a 45% ethanolic solution of 0.01 N NaOH, 5 min at 70 C; rinsing in water and followed by 1 min in a 1% aqueous solution of acridine orange made up in 0.02 N NaOH, also at 70 C, then washed, and dried on slides. Method II. 0.5% Janus green in aqueous 0.05 N NaOH, 5 min at 70 C; rinsing in water then into 0.5% Darrow red in 0.05 N NaOH (aq.), 2 min at 70 C., washing, and drying on slides. Method III. 1% methyl green (commercial, unpurified) in 1% aqueous borax for 15-20 min at 20-25 C, washing and attaching to slides. All staining was performed by floating the sections on the staining solutions, all drying at 70 C, and mounting in a resinous medium. T-AO gave blue to violet cytoplasmic structures, darker nuclei which contrasted strongly with yellow connective tissue and the secretion of goblet cells. JG-DR resembled a hematoxylineosin stain, but by shortening the staining time in DR to 0.5-1 min, collagenous and elastic tissue retained more of the green dye. MG-MV gave dark green nuclei in light green cytoplasm, with collagenous and elastic tissues in blue to violet. As with most methods for staining ultrathin sections, thicknesses of less than 1 μ required longer staining times.     

Staining Residual Lipids in Ultrathin Sections of Tissues Embedded in Polyester Resin

Rat suprarenal glands fixed in Palade's 1% OsO₄, buffered at pH 7.7 with veronal-acetate, to which 0.1% MgCl₂ was added, were embedded in Vestopal-W and sectioned at 0.2-1 µ. The sections were attached to slides by floating on water, without adhesive, and drying at 60-80° C, placed in acetone for 1 min and then treated with the following staining procedure: Place the preparation in a filtered solution of oil red O, 1 gm; 70% alcohol, 50 ml; and acetone, C.P., 50 ml; for 0.5-1 hr. Rinse in absolute ethyl alcohol; drain; counterstain with 0.5% aqueous thionin for 5 min; rinse in distilled water; drain; stain in 0.2% azure B in phosphate buffer at pH 9, for 5 min. Dry and apply a drop of immersion oil directly on the section. The preparations are temporary. Ciaccio-positive lipids, rendered insoluble by OsO, fixation, stained red to ochre.     

Simplified Aldehyde-Fuchsin Staining of Neurosecretory Cells

Gomori's original aldehyde-fuchsin method has been modified by the combination of Halmi's counter stain with Gabe's preparation, consisting of basic fuchsin, 1 gm; boiling water, 200 ml; with HC1, 2 ml and paraldehyde, 2 ml added after cooling and filtering. The solution so made was allowed to ripen 3-4 days at room temperature, and the precipitate which formed was filtered off and dried at 55-60°C. The staining solution consisted of 0.5 gm of the dry precipitate dissolved in 100 ml of 70% alcohol. The staining follows original procedures except that it is very important to bring slides from water to 70% alcohol before placing them in the aldehyde-fuchsin solution and also to remove all excess staining solution by rinsing in 95% alcohol after staining. The staining solution is stable for at least 6 mo.     

New Rapid Methods for Tissue Diagnosis

Two new methods applicable to the staining of fixed and fresh frozen tissue sections are presented herein. In addition certain improvements are suggested for the technic reported by Geschickter, Walker, Hjort and Moulton (1931). In brief the procedures are as follows:

1. The thionin eosinate method of Geschickter et al (1931). This procedure has been modified as follows:
   1. A mixture of diethylene glycol, 40 parts, ethylene glycol, 40 parts, and grain alcohol, 20 parts is superior to ethylene glycol, 80 parts, and ethyl alcohol, 20 parts, as a solvent for the compound stain in that the staining is intensified.
   2. Ethylene glycol monobutyl ether supplants diethylene glycol monobutyl ether because of its lower viscosity.
   3. Ethyl phthalate replaces butyl phthalate on account of a more satisfactory viscosity.
2. The methyl green eosinate procedure is the same as the modified thionin eosinate method except for the following variations:
   1. The staining time is increased to one minute.
   2. Decolorization and washing are reduced to about 15 seconds.
3. The hematoxylin-eosin method. After cutting, the tissue sections are carried thru the following steps:

Unfold in water; transfer to formalin (4 to 40%) for at least 30 seconds; stain in hematoxylin (Harris) for 30 to 60 seconds; wash in water, 5 seconds; decolorize in 0.1% HC1 or saturated aqueous picric acid, 5 seconds; wash in water, S seconds; float in 0.5% ammonia, 5 to 10 seconds; wash in water, 5 seconds; stain in 5% aqueous eosin, 15 seconds¹; wash in water, 5 to 10 seconds; dehydrate in a mixture of diethylene glycol, 30 parts, and ethyl alcohol, 70 parts, 5 to 10 seconds; dehydrate in ethylene glycol monobutyl ether, 5 to 10 seconds; clear in ethyl phthalate, 5 to 10 seconds; float on a glass slide, blot with photographic lintless blotter, place a drop of neutral gum damar on the section, and cover with glass cover slip.