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.     

Staining of Negri Bodies

The following procedure for staining Negri bodies in sections is based on methods previously described by MacNeal, by Haynes, and by Richter:

Fixation:

1. 1. Zenker's solution 4 hours at 37°C or Dominici's 3 hours.
2. 2. 70% alcohol, 12 to 18 hours at room temperature.
3. 3. 80% alcohol, about 5 to 6 hours.
4. 4. 90% alcohol, about 4 to 6 hours.
5. 5. Absolute alcohol about 16 hours.
6. 6. Ether and absolute alcohol aa, about 8 hours.
7. 7. 16 to 24 hours in the following mixture: celloidin 1 g., methyl salycilate 25 cc., abs. alcohol 25 cc., ether 25 cc.
8. 8. Chloroform and paraffin, 2 to 3 hours.
9. 10. Paraffin, 1 to 1 1/2 hours.
10. 11. Embed.

staining:

1. 1. Cut sections 4 to 5 μ.
2. 2. Bring section to water and cover with Lugol's iodine for 10 minutes.
3. 3. Decolorize with a 2% sodium thiosulfate (hypo).
4. 4. Wash thoroly with water.
5. 5. Cover with a mixture of equal parts of 0.5% phloxine and 1% eosin Y (National Aniline brand) and leave for 15 minutes.
6. 6. Wash with water and stain 2 to 5 minutes in 0.1% azure B (National Aniline).
7. 7. Wash with 96% alcohol and decolorize in a mixture of 2 parts absolute alcohol with 1 part clove oil, ordinarily for not more than 1/2 to 1 minute.
8. 8. Dehydrate rapidly, clear, and mount in Yucatan Elemi.

     

A Note on the Gram Stain

A modification of the Gram stain in which iodine-alcohol is substituted for 95% alcohol as a decolorizing agent has been found particularly useful in staining Gram-positive organisms in tissues and also for smears. The technic for tissue sections follows:

1. Apply nuclear stain.
2. Wash.
3. Stain in Hucker's gentian violet 2 to 3 minutes (i. e. 1 part Sat. Alc. Sol. crystal violet to 4 parts 1% Aqu. Sol. ammonium oxalte).
4. Wash in water.
5. Stain in Gram's iodine 5 minutes.
6. Wash in water.
7. Decolorize in 95% alcohol to which enough tincture of iodine has been added to give a mahogany color.
8. Counterstain.
9. Dehydrate and mount.

     

Method for Staining Bacterial Flagella

The following method of staining bacterial flagella is ecommended for use on smears made from suspensions of 10 to 16-tour agar slant cultures, incubated 30 minutes at 37°C before spreadng on thoroly cleaned and named slides:

1. Cover with fixative (100 cc. of 1/4 sat. aqu. solution picric acid, with 5 g. tannic acid and 7.5 g. ferrous sulfate).
2. Wash with tap water, dry and cover with Fontana spirochaete stain; heat to steaming and allow to act for 1 to 2 minutes. Wash in ap water. The stain is prepared as follows: To 25 cc. 2% AgNO₃ add dilute ammonia till the precipitate which forms redissolves; then add more AgNO₃ till a faint turbidity results. A clear solution is useess.

     

The Bielschowsky Staining Technic a Study of the Factors Influencing Its Specificity for Nerve FIBERS

By comparing results obtained with adult mammalian tissue from introducing variables into each separate step in block-staining by the Bielschowsky silver method, the following conclusions were reached:

1. No specific means for inhibiting the staining of connective tissue and still permitting complete staining of nerve fibers was found, but the avoidance of overstaining was very helpful toward such differentiation.
2. Overstaining could be corrected by reducing the concentration of the silver nitrate bath or by adding an excess of ammonia to the ammoniated silver bath.
3. Staining of fine fibers was favored by adding acetic acid to the formaldehyde used for fixation or by adding pyridin to the silver nitrate bath.
4. Addition of protein-precipitating organic acids (trichloracetic or sulfosalicylic) to the fixative was disadvantageous.
5. Prolonged fixation favored an increase in intensity of the stain. Four days' time was sufficient.
6. Extraction of lipids with ammoniated alcohol gave results similar to those obtained after extraction with pyridin, but the stain was lighter.
7. Ammoniated silver carbonate without excess ammonia had an action similar to ammoniated silver hydroxide with excess ammonia.
8. An excess of ammonia in the ammoniated silver solution (Ag 0.1 N) was tolerated, without apparent impairment of nerve-fiber staining, up to 6 M NH₃, altho the use of more than 3 M excess (2 cc. concentrated ammonia water added to 100 cc. of balanced ammoniated silver hydroxide solution) seemed unnecessary.
9. Impregnation with 1.7% (0.1 N) silver nitrate solution was quite satisfactory and variations in the concentrations of this bath suggested that the practical limits of concentrations that would be generally satisfactory lay between 0.3 and 3.0%.
10. The writers' experiences agreed with Agduhr's relative to the advantage of washing in 2.5% acetic acid between the ammoniated silver bath and formaldehyde reduction.

     

Toxic effects of some alcohol and ethylene glycol derivatives on Cladosporium resinae.

Eleven commercially available alcohol and ethylene glycol derivatives were tested for their toxicity toward a problem organism in jet fuel, Cladosporium resinae. In the presence of glucose, 20% (vol/vol) ethylene glycol monomethyl ether prevented spore germination and mycelial growth, and 10% (vol/vol) 2-ethoxybutanol, 10% 2-isopropoxyethanol, 10% 3-methoxybutanol, 5% 2-butyloxyethanol, 5% ethylene glycol dibutyl ether, and 5% diethylene glycol monobutyl ether were found to have similar effects. In a biphasic kerosene-water system, 3-methoxybutanol, 2-butyloxyethanol, and diethylene glycol monobutyl ether were again found to be more toxic than ethylene glycol monomethyl ether. Considerable potassium efflux, protein leakage, and inhibition of endogenous respiration were observed in the presence of the more toxic compounds. 2-Butyloxyethanol also caused loss of sterols from cells.     

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.     

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

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

Staining Paraffin Sections with Protargol: 1. Experiments with Bodian's Method. 2. Use of Jvpropyl and N-Butyl Alcohol in Hofker's Fixative:

Paraffin sections of nervous tissue, which had been fixed in Hofker's fluid, stained readily with protargol solution without the addition of metallic copper or other activator. Amidolsulfite mixtures reduced the protargol more rapidly and completely than hydroquinone-sulfite. Intensification of the stain could be secured by reducing with 0.5% amidol (or pyrogallol) solution after gold toning. The completeness of staining of unmyelinated fibers of the dorsal roots of cat spinal nerves was checked by estimating the number of fibers in a root and the cells of its associated ganglion. A fiber cell ratio of 1:1 was found hi 4 specimens, indicating within limits of error that all fibers were stained. An improvement of die original Hofker's mixture as a fixative was obtained by using a mixture of formic acid, 5 cc.; trichloracetic acid, 10 g.; n-propyl alcohol, 20 cc.; and n-butyl alcohol, 60 cc. (instead of the acetic, trichloracetic, ethyl alcohol mixture used hi the original formula). The following arbitrary method is suggested. Fix 12 to 24 hours, pass to water thru graded ethyl alcohol, wash several hours, dehydrate and embed in paraffin. Cut, mount, and remove the paraffin, pass to water and impregnate 2 or 3 days at 27 to 30$$C. in a 0.5% aqueous solution of protargol (Winthrop Chemical Co.). Rinse 2 or 3 seconds and reduce with 0.5% amidol (Agfa brand used) in 5% sodium sulfite solution. Wash, tone with 0.1% gold chloride, wash and reduce with 0.5% amidol (no sulfite), wash, dehydrate and cover. The method works well on spinal nerve roots, cerebrum, cerebellum, and spinal cord, and moderately well on nerve trunks including sympathetic nerves. Tissues from cat and guinea pig were used.     

Staining Paraffin Sections with Protargol 6. Impregnation and Differentiation of Nerve Fibers in Adrenal Glands of Mammals

A series of experiments with protargol staining of nerve fibers in mammalian adrenal glands has yielded the following procedure: Fix-1-2 days in a mixture of formamide (Eastman Kodak Company) 10 cc, chloral hydrate 5 g., and 50% ethyl alcohol 90 cc. Wash, dehydrate and embed in paraffin. Cut sections about 15 and mount on slides. Remove the paraffin and run down to distilled water. Mordant 1-2 days in a 1% aqueous solution of thallous (or lead) nitrate at 56-60°C. Wash thru several changes of distilled water and impregnate in 1% aqueous protargol (Winthrop Chemical Company) at 37-40°C. for 1 to 2 days. Rinse quickly in distilled water and differentiate 7-15 seconds in a 0.1% aqueous solution of oxalic acid. Rinse thru several changes of distilled water for a total time of 0.5 to 1.0 rain. Reduce 3-5 rain, in Bodian's reducer: hydroquinone 1 g., sodium sulfite 5 g., distilled water 100 cc. Wash in running water 3-5 min. and tone 5-10 min. in a 0.2% gold chloride solution. Wash 0.5 min. or more and reduce in a 2% oxalic acid solution to which has been added strong formalin, 1 cc. per 100. (Caution. This last reduction is critical and over-reduction can spoil an otherwise good stain; 15-30 seconds usually suffices, and the sections should show only the beginning of darkening to a purplish or gray color.) Wash, fix in hypo, wash, dehydrate and cover.     

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.     

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

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

On-The-Slide Impregnation by Albumen-OsO4, Selective for Ciliary Basal Bodies and Osmiophilic Granules in Synaptic Terminals

Procedure: Fix 24 hr by immersion in Heidenhain's Susa (2-4 mm specimens) or by perfusion for spinal cord or brain of cats or larger mammals. Wash in 80% alcohol containing 0.5% I₂, dehydrate, and embed in paraffin; or, better, double embed in celloidinparaffin. Attach sections to slides by albumen-glycerol. Remove paraffin, and celloidin if used, treat again with iodized alcohol for 30 min, followed by 0.25% Na₂S₂O₃, and wash well with distilled water. Impregnate in darkness for 5 days at 37 C in aqueous 0.66% OsO₄ to which 0.2% fresh egg albumen has been added. Check the impregnation microscopically and return the slide to the original staining solution for another 2-3 days if the granules do not show. Wash well in distilled water, dehydrate and cover as usual. The stain does not fade in water, alcohol or zylene; therefore almost any counterstain can be applied. The method stains selectively black the ciliary basal bodies and the osmiophilic granules in the majority of the different types of synaptic terminals; most red blood cells and a few nuclei also stain black.     

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.     

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.     

A Single solution iron-hematoxylin Stain for Intestinal Protozoa

A single solution iron-hematoxylin stain is described for staining fecal smears rapidly and simply. The stain is prepared from the following solutions: Solution A: 1% hematoxylin in 95% alcohol, prepared by diluting a stock solution of 10% hematoxylin in 95% alcohol. Solution B: Ferric ammonium sulfate (violet crystals), 4.0 g.; glacial acetic acid, 1.0 ml.; concentrated sulfuric acid (sp. gr. 1.8),0.12 ml.; distilled water, 100 ml. Mix equal parts of Solution A and Solution B; allow to stand overnight, filter and use. For maximum length of staining life, store in full, air-tight bottles. To stain fecal smears, fix in Schaudinn's, pass through iodine alcohol to 50% alcohol, stain for three minutes, wash in running tap water 5 to 15 minutes, dehydrate and mount.     

A Single solution iron-hematoxylin Stain for Intestinal Protozoa

A single solution iron-hematoxylin stain is described for staining fecal smears rapidly and simply. The stain is prepared from the following solutions: Solution A: 1% hematoxylin in 95% alcohol, prepared by diluting a stock solution of 10% hematoxylin in 95% alcohol. Solution B: Ferric ammonium sulfate (violet crystals), 4.0 g.; glacial acetic acid, 1.0 ml.; concentrated sulfuric acid (sp. gr. 1.8),0.12 ml.; distilled water, 100 ml. Mix equal parts of Solution A and Solution B; allow to stand overnight, filter and use. For maximum length of staining life, store in full, air-tight bottles. To stain fecal smears, fix in Schaudinn's, pass through iodine alcohol to 50% alcohol, stain for three minutes, wash in running tap water 5 to 15 minutes, dehydrate and mount.     

Glucose condensation by glucoamylase in organic solvents

Summary Oligosaccharides were synthesized through the enzymatic condensation of D-glucose by glucoamylase in water-organic mixtures with high concentrations of two of diethylene glycol diethyl ether or triethylene glycol dimethyl ether. The effect of water content on the yield of reaction was studied; maximum yield was obtained with 10% (v/v) of water in the two systems. Kinetics of synthesis and products composition were different with the two solvents. 37% of glucose were condensed by action of glucoamylase from a reaction medium containing 20 g/L of glucose and 90% (v/v) of diethylene glycol diethyl ether.     

Maxilon Blue Rl: A New Metachromatic Monoazo Dye

Maxilon blue RL, a basic monoazo dye developed by Geigy S. A., stains metachromatically acid mucopolysaccharide-containing elements in histological sections. This property is due to a chromatographically pure blue fraction which is the main component of the dye. The following routine has been developed for staining sections of formalin-calcium fixed paraffin-embedded tissues. Dewax in xylene and hydrate the sections through alcohol; stain in 0.05% aqueous Maxilon blue RL, from 30-60 sec; wash in distilled water; dehydrate either in tertiary butanol, 2 to 3 min, after removing excess water from the slide by blotting; or, rinse in 70% ethyl alcohol and dehydrate in 2 changes, 2 min each, of absolute alcohol; clear in xylene; mount in DPX or in Canada balsam. Acid mucopolysaccharides are colored red to violet; other basophilic elements, blue.     

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.