An Improved Hematoxylin-Eosin Stain for Sections of Marrow Units

The procedure recommended is: Fix “marrow units” (small functional structures of bone marrow) in 10% formol-saline solution for 1-2 hours and dehydrate in 80% alcohol, 95% alcohol and acetone 30 minutes each. Place in fresh 50° and 53°C. paraffin for 30 minutes each. Embed in fresh 53°C. paraffin. Serially section at 5μ thickness and mount with Schleicher's floating solution. Allow to dry for 1 hour in an oven and deparaffinize by passing through xylene I and II, absolute alcohol I and II, and 95% alcohol. Rinse in fresh distilled water and place in dilute Harris' hematoxylin (stock solution 50 ml., distilled water 200 ml.) for 2 to 3 minutes. Rinse well in distilled water and check staining under the microscope. Dip in acid-alcohol 5 times (1 dip to equal about 1 second). Rinse well in weak (0.02%) ammonia water and distilled water. Dip in 2% aqueous phosphotungstic acid about 3 to 5 times (equal to 3-5 seconds). Rinse in fresh distilled water and place in weak ammonia water for 1 minute. Rinse in fresh distilled water I and II. Place in 80% alcohol for 5 minutes and check under the microscope for “blueness” and nuclear differentiation. Place in dilute alcoholic eosin (0.5% alcohol-eosin stock solution 10 parts and 95% alcohol 90 parts) for 1 to 2 minutes. Rinse in 80% alcohol and place for 1 minute in 95% alcohol. Check under the microscope for staining quality. Place in absolute alcohol for 1 minute, alcohol-xylene (equal parts), 10 dips, and xylene I and II. Mount. This hematoxylin-eosin staining schedule brings out minute structural detail of bone marrow tissue heretofore not demonstrable.     

An Improved Pollen Grain Smear Method for Wheat

Anthers containing actively dividing pollen grains were treated 1 hour at 18-20° C. with 0.2% solution of colchicine, washed 1 hour in water, soaked in 0.002 M aqueous solution of 8-oxyquinoline at 10-14° C. for 1 hour, washed in water for 1 hour and then fixed in Carnoy's solution (alcohol, chloroform, acetic acid, 6:3:1) for 6 hours to overnight. They were washed successively in acetic-alcohol (1:1) 10-15 minutes, 70% alcohol 10-15 minutes and in water 30 minutes before hydrolysing them in bulk in 1 N HCl at 60° C. for 10-15 minutes. “Finally, they were stained in leuco-basic fuchsin for 15-30 minutes. Pollen grains were squeezed out of a stained anther in a small drop of egg albumen on a slide and the albumen smeared uniformly on the slide. The slide was dipped successively for a few seconds in glacial acetic acid and 45% acetic acid respectively. The smear was covered by a cover glass in a drop of aceto-carmine and pressed gently between folded filter papers. The cover glass was sealed with paraffin and stored overnight. To make the preparation permanent the paraffin was removed and the cover glass separated in a 1:1 mixture of acetic acid and n-butyl alcohol. The slide and the cover glass were then passed through n-butyl alcohol, 2 changes, and finally remounted in balsam.     

An Ammoniated Silver Carbonate Technic for Nervous Structures in Mounted Paraffin Sections

Specimens of both vertebrate and invertebrate nerve-containing tissues were fixed 2-3 days in Bouin's fluid, soaked 2 days in alcohol containing 2% strong ammonia water, dehydrated and embedded in paraffin. The sections were mounted with gelatin adhesive according to Masson's procedure, dewaxed, passed through graded alcohols to water, then back to 2% ammoniated 80% alcohol for 12-24 hours. The slides were rinsed 3-5 seconds in distilled water, impregnated about one and a half hours in 40% AgNO₃ at increasing temperature up to 45°C. The slides were flooded with 62.5% formalin and this solution allowed to remain 3-5 minutes; they were then blotted with filter paper. A second impregnation in ammoniated silver carbonate, controlled under the microscope, was followed by a 10-minute treatment with 10% aqueous acetic acid, toning with gold chloride, then thiosulfate and finally washing. Counterstaining with ponceau red or acid fuchsin, eventually followed by aniline blue or fast green, dehydration and covering, completed the process.     

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 new method for surface staining large slices of fixed brain using a copper phthalocyanine dye

Here we describe a method for gross staining of gray matter in slices of formaldehyde-fixed human brain. After protection of white matter with 4% phenol at 60°C for 5 min followed by a cold water wash, the gray matter was stained for 10-15 min at 20-25°C with 1% aqueous copper(II) phthalocyanine tetrasulfonic acid tetrasodium salt (CPTS). The staining resisted all attempts to be washed from the gray matter. Stained slices can be stored indefinitely in slightly acidified water, or plastinated as permanent dry specimens.     

A One-Solution Stain for Spermatozoa

Fresh semen is allowed to liquefy 30-60 minutes and thin, even smears of it made on clean slides or cover glasses. The smears are fixed 3 minutes with an equal-parts mixture of alcohol and ether, then air dried. They are stained 5-7 minutes in an aqueous solution made by mixing 2 volumes of 5% aniline blue (water soluble), 1 volume of 5% eosin B and 1 volume of 1% phenol. Staining at 40-60°C. is recommended. After staining, the smears are washed with distilled water, air dried and mounted in balsam or synthetic resin. The method was used on over 2000 samples of dog semen and some human specimens. Good preservation and differentiation of cytological structures was obtained uniformly, but tests were not made with other species.     

Combined Carbowax-Paraffin Technic for Microsectioning of Fixed Tissues

A combined Carbowax-paraffin technic for microsectioning fixed tissues gave ribbon sections as do paraffin infiltrated and embedded tissues. Blocks of formalin or alcohol fixed tissues 2 mm. thick were infiltrated with H.E.M. (Polyethlene Glycol: Carbowax, Hartman-Leddon Co.) or with one of the following polyethylene glycol ester waxes (Glycol Products Co., Inc.) for 4 hours at (61°C.): Polyethylene Glycol 600(Di) Stearate; Carbowax 1000-(Mono) Stearate; Carbowax 4000 (Mono) Stearate; Carbowax 4000 (Mono) Laurate; Carbowax 6000 (Mono) Oleate. The Carbowax infiltrated tissues were placed for 10 minutes in xylene (61° C.), into paraffin (61° C.) for 30 minutes, then into molten paraffin contained in separate molds. (The xylene passage can be excluded for preparations which preclude its use). The blocks were hardened rapidly by submerging in ice water and were fastened to carriers as in the usual paraffin technic. Tissues were cut 6 µ thick. Segments of ribbon were spread on a water bath and mounted on slides. After drying, tissues were stained directly with hematoxylin-eosin or were carried through xylene and alcohols as in routine paraffin preparations prior to staining. The Sudan III fat stain and Best's carmine stain for glycogen were applied as in usual technics. Cellular detail was well preserved and structures did not show the extent of distortion and shrinkage encountered in ordinary paraffin technic preparations.     

Superoxide Production by Stimulated Neutrophils: Temperature Effect

Activation of neutrophils results in a one-electron reduction of oxygen to produce the superoxide anion and other oxygen-derived, microbicidal species. Evidence from many kinetic studies of oxygen-derived radicals generated by stimulated neutrophils in vitro shows that radical production is optimal at 37°C but only lasts several minutes and then rapidly subsides. These findings support the widely held perception that the neutrophil's “oxidative burst” is a transitory event that peaks within minutes of stimulation and ends shortly thereafter. However, while some studies have shown that under controlled conditions stimulated neutrophils can generate superoxide continuously for several hours, others have observed that the superoxide formation by neutrophils stimulated in buffer at 37°C does not persist. To reconcile the conflicting findings and to better understand neutrophil function, we have reinvestigated the effect of temperature on the kinetics of radical generation by PMA-stimulated cells. Electron paramagnetic resonance spectroscopy coupled with spin-trapping and SOD-inhibitable ferricytochrome c reduction were used to monitor superoxide production by neutrophils stimulated at either 25°C or 37°C in RPMI 1640 medium or in Hank's balanced salt solution. When oxygen was supplied continuously, neutrophils stimulated at 25°Cin buffer or in medium generated superoxide for several hours but at 37°C. particularly in HBSS, O₂-formation strikingly and rapidly decreased. This cessation of superoxide generation was reversible by lowering the temperature back to 25°C. These data imply that in vivo neutrophils may be capable of generating oxy-radicals for prolonged periods. In part, our results may also explain the often observed termination of neutrophil-derived radical formation in vitro and help to dispel the perception that neutrophil-derived oxy-radical production is an ephemeral phenomenon.     

Locating Iodine in Tissues Autographically, Especially after Fixation by Freezing and Drying

The ability of radioactive elements to affect photographic emulsions enables the detection of radioactive iodine in the thyroid. By placing unstained histological sections of a thyroid (from an animal treated with radioactive iodine) in contact with the gelatin side of medium lantern slide plates, each accumulation of radioactive iodine in the section affects the photographic plate. After exposures prolonged for several days to several weeks depending on the amount of radioactivity in the tissues, the plate is developed and fixed by routine photographic methods. The histological section is stained and may be compared under the microscope to the reactions on the plate or “autographs”.

In an attempt to detect the location of the inorganic iodine which is displaced during fixation and embedding by ordinary methods because of its solubility, a simplified freezing-drying technic for fixation was devised which, at least with the thyroid, yielded well fixed sections. The quick freezing was obtained with acetone-dry-ice mixtures; and the drying was performed at -25° to -30° C. Preliminary addition of paraffin to the tube in which the drying was performed made possible the inclusion in vacuum by heating the tube when drying was completed. The tissue could then be sectioned at 10ju on the microtome. The slides were placed on photographic plates for detection of radioactive iodine as indicated above. Before staining, the sections were treated with absolute alcohol for denaturation of the proteins.     

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.     

Staining Nerve Fibers After Sublimate-Acetic and After Bouin'S Fluid

A silver staining method for paraffin sections of material fixed in HgCl₂, sat. aq., with 5% acetic acid is as follows. Process the sections through the usual sequence of reagents, and including I-KI in 70% alcohol, thiosulfate (5% aq.), washing and back to 70% alcohol containing 5% of NH₄OH (conc. aq.). After 3 minutes in the ammoniated alcohol, wash through tap water and 2 changes of distilled water and silver 5-10 minutes at 25°C. in 15% AgNO₃ aq. to which 0.02 ml. of pyridine per 100 ml. has been added. Blot the slide, but not the section and do not rinse. Reduce at 45°C. in 0.1% pyrogallol in 55% alcohol, then rinse in 55% alcohol and wash in water. The remainder of the process consists of gold toning, intensifying in oxalic acid, fixing in 5% Na₂S₂O₃, washing, dehydrating, clearing and covering. When the specimen contains much smooth muscle, the I-KI solution is acidified before use by adding 2 ml. of 1N nitric acid per 100 ml., and the sections treated for 3 minutes instead of the usual 2 minutes. Formalin should not be added to sublimate-acetic, but specimens that do not contain strongly argyrophilic nonneural tissue may be fixed in formalin or, preferably, Bouin's fluid. Sections of tissue after the latter type of fixation will not require the I-KI and thiosulfate but can go from 95% alcohol to the ammoniated alcohol. The advantages of fixing in HgCl₂-acetic acid are suppression of the staining of connective tissue and intensifying the staining of nerve fibers.     

Autonomic thermoregulatory effects of electrical stimulation of the hypothalamus in the sheep (Ovis aries)

1. 1.|The effects of electrical stimulation of the preoptic region, on autonomic thermoregulatory responses, were studied in conscious sheep at ambient temperatures of 5, 20, and 40°C.

2. 2.|Stimulation of the dorsal preoptic region elicited co-ordinated thermoregulatory responses characterized by increased respiratory frequency (RF), vasodilation of the ears and lowered body temperature. Stimulation inhibited shivering in cold environments.

3. 3.|The thermoregulatory responses were greater at 5°C in unshorn than in shorn sheep. Increased RF, induced at 20 and 40°C, persisted several minutes after stimulation ceased.

4. 4.|Intraventricular injection of noradrenaline reduced both normal and electrically-induced panting.

5. 5.|Sheep would press panels to electrically stimulate the preoptic region and this “self-stimulation” activated heat-loss mechanisms.

A Controllable Silver Stain for Nerve Fibers and Nerve Endings

A paraffin section method is described with a yellow-brown-black color range comparable to that of Ranson's pyridine silver block stain. After impregnation with activated protargol and reduction with a fine grain photographic developer, silver nitrate impregnation and reduction are repeated as often as necessary. The procedure is as follows:

Place hydrated sections of tissue fixed in chloral hydrate (25 g. in 100 ml. of 50% alcohol) in 1% aqueous protargol (Winthrop Chemical Co.) containing 5-6 g. metallic copper for 12-24 hours. After rinsing in 2 changes of distilled water, reduce 5 to 10 minutes in: Elon (Eastman Kodak Co.) 0.2 g., Na₂SO₃, dessicated, 10 g., hydroquinone 0.5 g., sodium borate powder 0.1 g., distilled water 100 ml. Wash thoroly in 4 or 5 changes of distilled water and place in 1% aqueous AgNO₃ for 10-20 minutes at 28°-50° C. Rinse in 2 or 3 changes of distilled water and reduce in the elon-hydroquinone solution. After thoroly washing in 4 or 5 changes of distilled water, examine under microscope.

If too pale, treat again in silver nitrate for 10-20 minutes, rinse, reduce 5-10 minutes and wash thoroly until nerve fibers show distinct microscopic differentiation, then dehydrate, clear and mount.     

Hydrolysis and Biodegradation of the Vesicant Agent HT: Two Potential Approaches

HT is a powerful vesicant produced for use as a chemical warfare agent. It is a mixture of 60 wt% 2,2' -dichlorodiethyl sulfide (“HD” or “sulfur mustard”) and 40 wt% bis-(2-(2-chloroethylthio)ethyl) ether (T). Because HT reacts with water to form primarily the alcoholic compounds thiodiglycol (TDG) and bis-(2-(2-hydroxyethylthio)ethyl) ether (T-OH), disposal might be accomplished by combining hydrolysis with biodegradation. The half-lives of H and T in a well-agitated 3.8% HT/water reaction at 90°C were 1.4 and 1.6 minutes, respectively. The concentrations of both compounds were reduced to less than 1 mg/L within about 30 minutes. TDG is readily biodegradable. However, T-OH biodegradability has not been reported previously. HBr treatment converted HT ether-alcohol products to TDG. A comparative study of two hydrolysis/biodegradation approaches is reported here. HT was hydrolyzed (1) in water, and (2) in water then with HBr. Products were used as feed for separate aerobic sequencing batch reactors (SBRs), and bioreactor performances were compared. Although both feed solutions were detoxified in the SBRs, water hydrolysis alone yielded better overall bioreactor operation, a more favorable mass balance, and a simpler process than with the HBr step. Results indicated that although the HBr converted ether-alcohol products to TDG, the HT products were biodegraded with greater efficiency when the HBr treatment was omitted.     

Mounting Frozen Sections with Gelatin

Frozen sections, 15-50 µ thick, are soaked for 5 minutes or longer in a mixture of equal parts of 1.5% aqueous gelatin and 80% alcohol, and teased onto a slide. After allowing excess fluid to evaporate, sections will be moist and can be blotted with filter paper that may require dampening with 95% alcohol. Immersed in 95% alcohol, the remaining gelatin will congeal, anchoring the section to the slide. If necessary, the sections can subsequently be coated with celloidin.     

A Urea Silver Nitrate Method for Nerve Fibers and Nerve Endings

A silver nitrate stain for nerve fibers and endings applicable to paraffin sections on the slide utilizes the properties of urea to accelerate the procedure and improve the specificity of the stain. After removal of the paraffin the sections are run through absolute, 95% and 80% alcohol and placed for 60-90 minutes at 50-60°C. in: 1% aqueous silver nitrate, 100 ml.; urea, 20-30 g.; 1g. mercuric cyanide and 1 g. picric acid in 100 ml. of distilled water, 1-3 drops. After the silver bath they are rinsed quickly in 2 changes of distilled water and reduced for 3-5 minutes at 25-30°C. in: water, 100 ml.; sodium sulfite, anhydrous, 10g.; hydroquinone, 1-2g.; urea, 20-30g. They are then washed thoroughly in 4-5 changes of distilled water, passed through graded alcohols into 80% alcohol and examined under the microscope. If nerve fibers are not distinct, the sections are returned to the same urea-silver-nitrate bath for 10-15 minutes, rinsed, reduced, washed and dehydrated as before. This process may be repeated until staining is adequate; then they are dehydrated, cleared, and mounted.

Nerve fibers show a color range from brown to black; nerve cells from yellow to brown; and the background, depending on the type of tissue and its fixation, from yellow to light brown.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

An Aceto-Osmium Method for Demonstrating Nematodes in Citrus Roots

Difficulties are encountered in observing nematodes in citrus feeder roots because of the presence of suberin and other unsaturated compounds. To obviate these difficulties, infected citrus roots, either fresh or preserved, are immersed in a covered jar for 2 hr at 52° C in a solution composed of distilled water, 16 parts; 10% acetic acid, 10 parts; and 2% aqueous osmium tetroxide, 2 parts. The stained, blackened roots are washed in running water for at least 1 hr and then bleached in 10-30% hydrogen peroxide at 32°C for a few seconds until the color of the roots lightens perceptibly. After several washings in water to stop the oxidation reaction, roots are dehydrated in 70, 95, and absolute ethanol held at 52 °C for 30 min at each concentration. After dehydration, roots are cleared in methyl salicylate at 52°C. Examination for nematodes in most cases, can be made after 30 min.     

Staining and Mounting Helminths

The following procedure is recommended: Fix ces-todes and trematodes (while held flat between glass slides) 0.5-2.0 hr. in the following mixture: formalin, 15; acetic acid (gl.), 5; glycerol, 10; 95% ethyl alcohol, 24; distilled H₂O, 46; all proportions by volume. After freeing them from the slides, wash thoroughly in running water and stain immediately thereafter. Stock staining solution: ferric ammonium alum (violet cryst.), 2 g.; distilled H₂O (cold) 100 ml.; after solution, add 2 ml. concentrated H₂SO₄, bring to a boil; add 1 g. coelestin blue B (Nat. Aniline), boil 3-5 min.; cool and add 10 ml. absolute methyl alcohol and 10 ml. glycerol. Dilute 1 vol. with 3 vol. distilled H20 for use. Stain 5-30 min., depending on size of specimens. Wash with 2 changes 0.5 hr. each of distilled H₂O, then 50% isopropyl alcohol 12-16 hr., 50% isopropyl alcohol 2 hr., followed by graded isopropyl alcohol for dehydration. Ether: ethyl alcohol (equal parts), 1 hr., is followed by embedding in celloidin in a sheet just thick enough to cover the specimens. Trim embedded specimens and dehydrate with isopropyl alcohol, 80%, 90% and absolute. Clear in beechwood creosote. Mount in balsam with cover glasses that overlap the edges of the celloidin 1-2 mm. While drying at 37°C, refill edges of mount with fresh balsam as needed. When dry, remove excess balsam and ring the edges with ordinary gloss enamel paint.     

Nile Blue Histochemical Method for Phospholipids

The technic recommended is: Fix 6-12 hr. in 10% formalin containing 1% CaCl₂. Cut frozen sections without embedding or after gelatin or carbowax. Stain 90 min. at 60°C. in saturated aqueous Nile blue sulfate, 500 ml. plus 50 ml. of 0.5% H₂SO₄, boiled 2 hr. before use. Rinse in distilled water, and place in acetone heated to 50°C. Remove the acetone from the source of heat and allow the sections to remain 30 min. Differentiate in 5% acetic acid 30 min., rinse in distilled water, and refine the differentiation in 0.5% HCl for 3 min. Wash in several changes of distilled water and mount in glycerol jelly. Results: phospholipids - blue; everything else - unstained. Counterstaining nuclei with safranin is optional, but if done, it preferably precedes the Nile blue and is then differentiated by the acetic acid. The histochemical principles on which the method is based are as follows: (1) The calcium compounds of phospholipids combine with the oxazine form of Nile blue sulfate and survive subsequent treatment; (2) neutral lipids are dissolved out by acetone; (3) proteins and other interfering substances are destained by the acetic acid and hydrochloric acid baths.