A Rapid Technic for Demonstrating mast cells in Mouse Skin

A simple rapid technic is described for demonstrating mast cells in mouse skin. The procedure requires about 60 minutes from time of specimen removal until permanent mounting. The steps comprise: (1) stretch-mounting of skin on a cardboard frame; (2) fixing and dehydrating in absolute ethanol for 15 minutes; (3) xylene washing for 10 minutes; (4) absolute ethanol washing for 10-15 minutes; (5) 3-4 minutes in a 0.1% aqueous solution of methylene blue; (6) dehydrating and differentiating in absolute ethanol; (7) clearing in xylene; (8) trimming and mounting. Cell counts may be made immediately, as well as high dry and oil immersion study of cytological detail of mast cells.     

Alcoholic Thionin Counterstaining Following Golgi Dichromate-Silver Impregnation

Brains of cats that had been fixed 2 months or longer in 10% formalin were cut into 3-6 mm. slices and impregnated by Golgi's dichromate-silver procedure (6% dichromate solution, 4-6 days; 1.5% silver nitrate solution 2 days). Sections 100 µ thick were cut after embedding in low melting point paraffin. Three changes of xylene and three of absolute alcohol were followed by staining 3-5 minutes in a saturated solution of thionin in absolute alcohol. The sections were dipped quickly in absolute alcohol and cleared in xylene, then differentiation was effected by an equal-parts mixture of absolute alcohol and xylene. A final clearing in three changes of xylene and mounting in Permount completed the process. Counter-staining was most successful when applied to freshly cut sections.     

An Analysis of Methods for Permanently Mounting Ovules Cleared in Four-and-a-Half Type Clearing Fluids

Ovules cleared in benzyl benzoate-4 1/2 clearing fluid can be permanently mounted in Piccolyte or Permount by replacing the clearing fluid with absolute ethanol, upgrading the ovules in mixtures of ethanol and xylene (3:1, 2:2, 1:3, and xylene), and mounting them in either mountant under the supported coverglass of a Raj slide. Optical sagittal sections through the ovules resemble microtome sections in that the protoplasts are slightly shrunken away from the cell walls. The artifact is common in permanently mounted sections; fixation and paraffin infiltration are usually cited as the causes—its appearance in the whole-mounted ovules is caused by xylene. Although miscible with the clearing fluid, Euparal is the least satisfactory of the standard mountants for permanent preparations of cleared ovules and is best used with an equal quantity of clearing fluid for semipermanent preparations. A large quantity of Euparal in the mountant produces pronounced shrinkage. A method for permanently mounting cleared ovules with the clearing image unaltered employs a mountant which contains the ingredients of Spurr low viscosity embedding medium. Vinylcyclohexene dioxide (10 drops) is combined with diglycidyl ether of polypropylglycol (6 drops) and nonenyl succinic anhydride (26 drops). Ovules treated for 24 hr in benzyl benzoate-4 1/2 clearing fluid are passed through a graded series of clearing fluid-epoxy medium mixtures (3:1, 2:2, 1:3, and pure epoxy medium) at intervals of 15 minutes. One drop of dimethylaminoethanol, the cure accelerator, is then added to the epoxy medium and the ovules are mounted and covered immediately on a Raj slide. The preparation is cured in an oven at 60 C for 24 hr and observed with phase contrast or Nomarski interference optics.     

A HYDROXYL-DEPENDENT SILVER METHOD FOR NERVE AXOPLASM

Axoplasm is selectively impregnated by the following steps: (1) fixation in 10% formalin or in 10% formalin with added sucrose, 15%, and concentrated NH₄OH, 1%, for 1-7 days; (2) frozen sections; (3) extraction of the sections in 95% ethyl alcohol, absolute alcohol, xylene, and 95% ethyl alcohol and absolute alcohol, 1 hr each; (4) distilled water, 3 changes of 10 min each; (5) 20% AgNO₃ (aq.) at 25°C, 30 min; (6) distilled water, 3 changes of 1-2 sec each; (7) 6.9% K₂CO₃, 1 hr; (8) water, 3 changes of about 1 min each; (9) 0.2%AuCl₃, 2 min; (10) distilled water; (11) 5% Na₂S₂O₃, 2 min; (12) washing, clearing and mounting. This procedure is proposed as a simplified stain for axoplasm, with other tissue components remaining unstained. The few reagents necessary suit this method for histochemical investigation of the mechanism of silver staining.     

A New and Rapid Method for Making Permanent Aceto-Carmin Smears

Smear the pollen mother cells of a single anther from each flower bud on a clean dry slide, using a small scalpel. Flood the slide with Belling's acetocarmin and heat for a second over an alcohol flame. Examine under the microscope to determine the stage of microsporogenesis. If the stage is satisfactory, smear the remaining anthers in the same manner, but fix and stain them by immediate immersion, face downward, in a petri dish full of hot (steaming) acetocarmin for from 1 to 10 minutes. Then rapidly transfer thru the following mixtures: two parts 99% (glacial) acetic acid plus one part absolute ethyl alcohol; one part acetic acid plus two parts absolute alcohol; and finally one part acetic acid plus nine parts absolute alcohol. The slides are then to be dehydrated completely by 1 to 2 minutes immersion in pure absolute alcohol, and cleared 2 to 3 minutes in a mixture of xylene and absolute alcohol in equal parts. The preparations are then made permanent by mounting each with balsam and a cover glass. The whole process takes from 5 to 15 minutes and is particularly recommended for chromosome counts.     

The Dehydration of Methylene Blue Stained Material Without Loss of Dye

A method is given for dehydrating methylene blue stained protozoan smears which should be applicable to the dehydration of tissues stained intra vitam with methylene blue. The procedure is: Wash with distilled water, place in tertiary butyl alcohol for 1 to 2 minutes, then in three or more changes of tertiary butyl alcohol for 15 minutes to an hour each, and mount directly in balsam or pass thru two changes of xylene before mounting.     

An analysis of methods for permanently mounting ovules cleared in four-and-a-half type clearing fluids

Ovules cleared in benzyl benzoate-4 1/2 clearing fluid can be permanently mounted in Piccolyte or Permount by replacing the cleaning fluid with absolute ethanol, upgrading the ovules in mixtures of ethanol and xylene (3:1, 2:2, 1:3, and xylene), and mounting them in either mountant under the supported coverglass of a Raj slide. Optical saggittal sections through the ovules resemble microtome sections in that the protoplasts are slightly shrunken away from the cell walls. The artifact is common in permanently mounted sections; fixation and paraffin infiltration are usually cited as the causes--its appearance in the whole-mounted ovules is caused by xylene. Although miscible with the clearing fluid, Euparal is the least satisfactory of the standard mountants for permanent preparations of cleared ovules and is best used with an equal quantity of clearing fluid for semipermanent preparations. A large quantity of Euparal in the mountant produces pronounced shrinkage. A method for permanently mounting cleared ovules with the clearing image unaltered employs a mountant which contains the ingredients of Spurr low viscosity embedding medium. Vinylcyclohexene dioxide (10 drops) is combined with diglycidyl ether of polypropylglycol (6 drops) and nonenyl succinic anhydride (26 drops). Ovules treated for 24 hr in benzyl benzoate-4 1/2 clearing fluid are passed through a graded series of clearing fluid-epoxy medium mixtures (3:1, 2:2, 1:3, and pure epoxy medium) at intervals of 14 minutes. One drop of dimethylaminoethanol, the cure accelerator, is then added to the epoxy medium and the ovules are mounted and covered immediately on a Raj slide. The preparation is cured in an oven at 60 C for 24 hr and observed with phase contrast or Nomarski interference optics.     

Histochemical Demonstration of Carbonic Anhydrase Activity

Fresh tissue slices fixed in chilled acetone for 1 hour and washed in distilled water for 10-30 minutes were incubated for 30-45 minutes at 37°C. in the freshly prepared incubating mixture: filtrate of a mixture of 8% sodium bicarbonate, 100 ml., and MnCl₂·4H₂O, 1 g. After washing in distilled water for 1 hour, they were dehydrated and embedded in paraffin. Sections were cut 15-20μU, deparaffinized, rinsed in absolute alcohol and placed in a 0.1% solution of potassium periodate for 48 hours at 37°C. The mounted sections were counterstained (if desired), dehydrated in alcohol, cleared in xylene (not carbol-xylene) and mounted in balsam. Many brown granules were produced on the sites of enzyme activity by this procedure. The results obtained seem to be in good agreement with previous findings by biochemical determinations.     

An Arginine Histochemical Method Using Sakaguchi's New Reagent

A mounted paraffin section of material fixed in Bouin's, Carnoy's or 10% formalin is allowed to stand 15 minutes at room temperature in a 0.3% solution of 8-hydroxyquinoline in 30% ethanol. The slide, with adhering solution, is placed in 0.15 N hypochlorite (with enough KOH added to make the solution 0.015 N KOH) for 60 seconds, then (without draining) into a solution containing: 10 ml. of 0.15 N KOH; 15 g. of urea; 70 ml. of tertiary butyl alcohol, and water to make 100 ml. Here it is gently agitated for 10 sec. and then kept in a second change of the same solution for 2 min. Two changes of pure tertiary butyl alcohol, 10 sec. and 4 min.; one in aniline, 3 min.; and one of 10 sec. in xylene, complete the procedure. Permount containing 0.02% aniline is used as a mounting medium.     

Hematoxylin-Eosin Staining of OsO4-Fixed Epon- Embedded Tissue; Prestaining Oxidation by Acidified H2O2

Successful application of hematoxylin-eosin staining to 0.5-1 μ sections of OsO₄-fixed Epon-embedded mammalian tissue is made possible by first treating the sections for approximately 1 min at 25-30 C with 10% H₂O₂ acidified with 0.1 or 0.01 N H₂SO₄ to pH 3.2. Subsequent steps are: washing; drying; Hams hematoxylin at 50 C, 1-2 min; washing; drying; 0.2-0.3% NH₄OH in 70% ethanol, 3-5 sec, drying at 50 C; 5% aqueous eosin for 3 & 45 sec at 25-30 C, washing; drying; clearing in xylene and mounting in resin. The use of acidified H₂O₂ prevents the staining of Epon and permits the characteristic staining picture to be obtained. Sections were attached to glass slides without adhesive and processed horizontally on a rack. Slides should be well drained and blotted before each drying step, to prevent formation of precipitate on the section.     

A Six-Dye Staining Schedule for Sections of Mesquite and Other Desert Plants

Materials are fixed in FPA (formalin, 2; propionic acid, 1; 70% ethanol, 17). Paraffin sections on slides are brought to 50% ethanol and stained as follows: (1) in Bismarck brown Y, a 0.02% solution in 0.1% aqueous phenol, 10-30 min; wash 30 sec in 0.7% acetic acid, and wash in distilled water 20-30 sec; (2) in crystal violet, 1% in 70% ethanol alkalinized with 1 drop of 1 N NaOH per 100 ml, 12-35 min; wash 30-60 sec in tap water to remove excess stain, and rinse 0.5 sec in 70% ethanol; then mordant in I₂-KI, 1% each in 70% ethanol, 40 sec, and rinse in 70% ethanol 2-5 sec; (3) in a mixture containing 0.4% acid fuchsin and 0.6% crythrosin B in 70% ethanol about 0.5 sec; rinse in 70% ethanol 5-15 sec to remove excess red; dehydrate in 70%, 95%, and absolute ethanol, 2-3 sec each; (4) in fast green FCF, 0.5% in a mixture of equal parts of methyl cellosolve, absolute ethanol, and clove oil, 5-15 sec; rinse in a mixture of clove oil, 10 ml; absolute ethanol, 100 ml; and methyl cellosolve, 10 ml, 5-7 sec; (5) in orange G, 0.75 gm in a mixture of clove oil, 40 ml; absolute ethanol, 40 ml; and methyl cellosolve, 60 ml, 5-30 sec; rinse clean in a 1:1 mixture of xylene and absolute ethanol, 5-20 sec Complete the clearing in pure xylene, 3 changes, 1.5 min in each, and apply a cover glass with synthetic resin. Slides are agitated in all steps except Bismark brown Y, crystal violet, and the xylenes. Contrast and staining intensity are adjusted by varying staining times in the dye solutions.     

Selective Silver Impregnation of Degenerating Axons In The Central Nervous System

Rat and rabbit brains containing surgical lesions of 5-10 days' duration were fixed in 10% formalin (neutralized with calcium carbonate) for 1 week to 6 months. Frozen sections (15-20 n) were rinsed and then soaked 7 minutes in a 1.7% solution of strong ammonia in distilled water. Subsequent treatment was as follows: rinse; 0.05% aqueous potassium permanganate 5-15 minutes; 0.5% aqueous potassium metabisulfite, 2 changes of 2.5 minutes each; wash thoroughly in 3 changes distilled water; 1.5% aqueous silver nitrate, 0.5-1.0 hr.; 1% citric acid, 5-10 sec.; 2 changes distilled water; 1% sodium thiosulfate, 30 see.; 3 changes distilled water. Each section is then processed separately. Ammoniacal silver solution (450 mg. silver nitrate in 10 ml. distilled water; add 5 ml. ethanol; let cool to room temperature; add 1 ml. strong ammonia water and 0.9 ml. of 2.5% aqueous sodium hydroxide), 0.5-1.0 min. with gentle agitation. Reduction of about 1 minute is accomplished in: distilled water, 45 ml.; ethanol, 5 ml.; 10% formalin, 1.5 ml.; 1% citric acid, 1.5 ml. Rinsing; 1% sodium thiosulfate, 10 sec.; thorough washing followed by dehydration through graded alcohol and 3 changes of xylene or toluene complete the staining process. Normal nerve fibers are slightly stained to unstained, degenerating fibers, black. The treatment in potassium permanganate is critical since too little favors overstaining of normal fibers and too much abolishes staining of degenerating fibers.     

The Use of Ba(OH)2 in Methanol for Demonstration of the Distribution of Sucrose in Sugarbeet Roots

Sucrose in the tissues of the sugarbeet (Beta vulgaris L.) can be shown as follows. Fresh or stored roots are cut into pieces having a block face of about 1 × 2 cm, and sections of about 150 μ thickness prepared from these. The sections are rinsed 15-30 sec in iced distilled water, placed in Ba(OH)₂-saturated methanol for 3 min and then rinsed twice in methanol for 1 and 5 min respectively. They are then transferred to ethanol through a graded series consisting of 80, 60, 40, and 20% methanol in ethanol, 5 min in each, with evacuation as necessary to remove bubbles. Temporary mounting is in ethanol and examination made by incident light or darkfield illumination. Gradual replacement of ethanol with xylene permits mounting in a resinous medium. An opaque granular precipitate of barium saccharate shows the location of the sucrose within the cells.     

Orthochromatic, Species-Limited Staining of Mast Cells With Night Blue

Night blue will stain the mast cells of rat, mouse and hamster selectively if alcohol differentiation is controlled. The technical steps are: Dewax paraffin sections with xylene, 2 changes; air dry; 2% Na₂SO₄, 3-5 sec; 0.5% night blue in 10% ethanol, 1 hr at 60°C; rinse in water; 9% HNO₃, 15 sec; water 1-5 min; 70% ethanol, 2 changes, 30 sec each; wash; 0.01% safranin, 3-5 sec; rinse, blot, air dry, mount in synthetic resin. A clear orthochromatic stain of the mast-cell granules occurs. Acid fixation prevents the staining reaction.     

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.     

Thick Sections for Embryology

Night blue will stain the mast cells of rat, mouse and hamster selectively if alcohol differentiation is controlled. The technical steps are: Dewax paraffin sections with xylene, 2 changes; air dry; 2% Na₂SO₄, 3-5 sec; 0.5% night blue in 10% ethanol, 1 hr at 60°C; rinse in water; 9% HNO₃, 15 sec; water 1-5 min; 70% ethanol, 2 changes, 30 sec each; wash; 0.01% safranin, 3-5 sec; rinse, blot, air dry, mount in synthetic resin. A clear orthochromatic stain of the mast-cell granules occurs. Acid fixation prevents the staining reaction.     

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.     

Hematoxylin Staining of Tissues Embedded in Epoxy Resins

Sections of 0.5-2 μ thickness are affixed to slides with albumen adhesive, thoroughly dried, and placed in xylene or toluene for 1 hr, then brought through ethanol to water. Sections of tissue fixed in O_sO₄ are treated first in 0.1% KMnO₄, then with 1.0% oxalic acid, and after rinsing, incubated at 60 C for 12-24 hr in hematoxylin (Harris's or Ehrlich's) and counterstained 10-15 min with 0.5% phloxine B. Permanent preparations are made by clearing and mounting in a synthetic resin. The method requires only easily available reagents and is suitable for routine processing of epoxy sections.     

Demonstration of Degenerating Nerve Fibers by a Modified Cajal Technic

Whole brains of cat were fixed in two changes of cold acetone (24 hours each) and embedded directly in paraffin. The degeneration time recommended is 5 days. Mounted sections 15-20 μ thick were deparaffined, washed in absolute alcohol and given successive treatments of 6 hours each with 1% ammoniated absolute alcohol and pure pyridine, washing well with distilled water between them and after the pyridine. Impregnation in 2% silver nitrate 12 hours at 30°C., rinsing in absolute alcohol and reducing in a 95% alcoholic solution of pyrogallol and formalin (3% and 5%) was followed by 50% alcohol, thorough washing in distilled water, toning in 1% gold chloride and intensification in 1% oxalic acid. Treatment in 10% sodium thiosulfate solution, washing, dehydrating and covering completed the procedure. Normal fibers, degenerating fibers and terminals were stained specifically.