Notes on Technic: A Device for Centering Tissue Blocks on the Porter-Blum Microtome

A method of double embedding fixed tissues in 3% low viscosity nitrocellulose and paraffin is described. Five percent phenol in 80% alcohol during dehydration and 5% glycerin in the nitrocellulose solutions enhance cutting qualities. A modified Ruyter's solution is used to flatten sections. After a section is aflixed to a slide, it is passed through chloroform and acetone to remove the paraftin and celloidin. A 1% celloidin dip insures adherence of the seaion to the slide. Slides are stored in 70% alcohol until they are to be stained. Following staining and dehydration in graded alcohols, clearing should be done in a 1: 3 mixture of terpineol and toluene.     

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.

     

Double Embedding Again

A method of double embedding fixed tissues in 3% low viscosity nitrocellulose and paraffin is described. Five percent phenol in 80% alcohol during dehydration and 5% glycerin in the nitrocellulose solutions enhance cutting qualities. A modified Ruyter's solution is used to flatten sections. After a section is aflixed to a slide, it is passed through chloroform and acetone to remove the paraftin and celloidin. A 1% celloidin dip insures adherence of the seaion to the slide. Slides are stored in 70% alcohol until they are to be stained. Following staining and dehydration in graded alcohols, clearing should be done in a 1: 3 mixture of terpineol and toluene.     

A Method for Injecting Insect Tracheae Permanently

By the use of an alcohol insoluble dye (trypan blue), acetic acid, and a detergent (“Santomerse No. 3”), a resulting dye solution is obtained which will completely penetrate the tracheal system of an insect. The dye is injected by the use of a vacuum and by the pressure produced when the air is allowed to re-enter the dye vessel. The dye itself is permanently fixed in the tracheae by means of a fixing solution containing alcohol, acetic acid and barium chloride as its components. The material must be properly preserved after staining. It may be stored indefinitely in 70% alcohol, xylol, cedar oil or clove oil, depending on whether the material is to be used for sectioning or for whole mounts. Injected material may be sectioned in either celloidin or paraffin, or may be cleared and mounted in toto.     

Protective Paraffin Infiltration of Soft Tissues in Insects to Facilitate Softening of Hard Exoskeletons

This technique can produce serial sections as thin as 5 μ from hard chitin-covered materials of insects or other arthropods. Procedures: Fix with alcoholic Bouin's fluid for 3 hr. Henceforth subject material to partial vacuum in each step to ensure a final proper embedding. Wash with 80% ethanol 2 or 3 times for 2 hr or until the picric acid is largely removed. Dehydrate to 90% ethanol and give 2 changes of n-butanol 2 hr each, and one of a 1:1 n-butanol-paraffin mixture in 56-57° oven for 12 hr. Finally, use 2 baths of pure paraffin, 3 hr each, to complete the infiltration. After the last bath, withdraw the specimen from the paraffin, and remove the superficial paraffin, first mechanically and then with a xylene bath for 4 min. Rinse first with n-butanol, and afterwards with absolute ethanol, 2 min each. The compound eyes are protected with a paraffin covering, the specimen is hydrated with a 1% aqueous solution of detergent for 1 hr and then washed with running tap water. The material is treated with a concentrated sulfuric-nitric mixture (H₂SO₄:HNO₃) for 4 hr to eliminate the exoskeleton. After this treatment, the specimen is washed with running tap water for 12 hr, dehydrated with acetone and then bathed in a 2% solution of celloidin in ethyl acetate to form a protective artificial cuticle. This coating is hardened with 2 quick baths of chloroform, the specimen reembedded in paraffin, and the block cast for sectioning.     

Basic Dye Counterstaining of Sections Impregnated by the Golgi-Cox Method

Celloidin blocks of Golgi-Cox impregnated material are cut at 50 μ, the sections collected in 70% alcohol, transferred to a 3:1 mixture of absolute alcohol and chloroform for 2 min, and then stored in xylene or toluene for at least 3 min, or up to 2 wk until processed further. Mounting is done on glass slides which have been coated with fresh egg albumen diluted in 0.2% ammonia water (or a 0.5% solution of dry powdered egg albumen) and then dried at 60°C overnight. For attachment to these coated slides, sections are first soaked for 2-3 min in a freshly prepared mixture of methyl benzoate, 50 ml; benzyl alcohol, 200 ml; chloroform, 150 ml; and then transferred quickly to the slides by means of a brush. After 2-3 min the chloroform evaporates and the celloidin softens. The slides are then immersed in toluene which hardens the celloidin and anchors the sections to the slides. Alcohols of descending concentrations to 40% are followed by alkalinizations, first in: absolute alcohol, 40 ml; strong ammonia water 60 ml, for 2 min, then in: absolute alcohol, 70 ml; strong ammonia water, 30 ml, for 1 hr. Excess alkali is then removed by 70% and 40% alcohol, 2 min each, and a 10 min wash in running tap water. Bleaching in 1% Na₂S₂O₃, for 10 min and washing again in tap water for 10 min completes the process preliminary to staining. The preparations are then stained for 90 min in an aqueous solution of either 0.5% cresylecht violet, neutral red, or Darrow red, buffered at pH 3.6. Dehydration and differentiation in ascending grades of alcohol, clearing with toluene or xylene, and applying a cover glass with a mounting medium having a refractive index of about 1.61 completes the process.     

A method for obtaining histological sections from tissue previously studied by scanning electron microscopy

An accelerated method of paraffin embedding of tissue specimens previously examined with scanning electron microscopy is proposed aimed to obtain sections for routine histological examination. The tissue is passed through acetone, absolute alcohol, alcoholic-oil celloidin solution, chloroform to be eventually mounted into paraffin. The method allows obtaining good quality sections within 24 hours.     

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.     

A Staining Combination for Phloem and Contiguous Tissues

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

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.     

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.     

Histological Localization of Microelectrode Placement in Brain by Ferrocyanide and Silver Staining

After recordings had been taken from a microelectrode used for mapping nerve impulses, a current of 100 μa from the positive pole of a direct current generator was run through the electrode for 5 sec while it was still in place. On terminating the experiment, in which the use of several electrodes was possible, 50-75 ml of a 1:1 mixture of 4% potassium ferrocyanide and 4% acetic acid was injected into each common carotid artery, and the brain left in situ for 0.5 hr. It was then removed and the electrode-bearing part fixed 5-6 hr in a 1:1 mixture of 40% formalin and 95% ethyl alcohol at 55 °C. This specimen was washed in running water 5-10 min, the electrodes removed and frozen sections of 40-80 μ cut and placed in 95% alcohol. Sections were stained 5-10 min at 25-30°C in 10% silver nitrate solution in 75-80% alcohol acidified by 3-4 drops of glacial acetic acid per 50 ml, washed 4-5 sec in each of 2 baths of 95% alcohol, and reduced while being agitated constantly in a 2% solution of pyrogallol and 6-7% formalin in 75-80% alcohol. Washing in 95% alcohol, clearing in clove oil or methyl salicylate followed by xylene and mounting in synthetic resin or balsam completed the process. Sites of electrolysis at the tips of electrodes (under magnification) were blue before silver staining and black after staining. Axons stained brown to black on a yellow background.     

The Fixation of Tissues Vitally Stained with Trypan Blue

The following fixative is recommended for tissues vitally stained with trypan blue: Chloroform, 2 parts; absolute ethyl alcohol, 2 parts; glacial acetic acid, 1 part; mercuric chloride to the point of saturation.

The tissue should be fixed 1 to 2 hours; transferred to 95% ethyl alcohol for 12 hours; to absolute alcohol for 12 to 24 hours; to a mixture of absolute alcohol and xylol for 1/2 hour, and finally to xylol, before embedding in paraffin. Cedar oil may be used for clearing in the place of xylol; in that case the tissues should be transferred from absolute alcohol to a mixture of absolute alcohol and cedar oil for 24 hours before placing in cedar oil alone.

Various counterstains can be used; Mayer's carmalum is excellent.     

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.     

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.     

Staining of Oligodendroglia with Silver Ammino Tungstate in Unembedded and Embedded Material

Specimens of brain or spinal cord fixed in formalin, Cajal's formol-bromide, or Koenig, Groat and Windle's formalin-acacia can be used to stain oligodendrocytes in frozen, in paraffin, or in celloidin sections. The sections are soaked 3-5 min in 0.02% acetic acid, pH 3.4, then rinsed 2-3 sec in 3% H₂O₂ and transferred to a silver bath prepared as follows: Mix equal parts of 10% AgNO₃ and 10% Na₂WO₄, and dissolve the precipitate with concentrated NH₄OH; avoid an excess of ammonia. Silver at room temperature for 15-20 sec, develop in 1% formalin, dehydrate, and mount. For embedded material, prepare a mixture consisting of 1 part of 10% aqueous Aerosol MA and 4 parts of 10% Aerosol OT in 95% alcohol. Add 5 drops of this mixture to each 50 ml of dilute acetic acid and 3% H₂O₂; 5 drops to each 20 ml of the silver bath.     

A Rapid Method for Preparing Paraffin Serial Sections of Teeth and Their Supporting Structures in the Dog

A method of preparing dogs' teeth for serial sectioning involving the following steps is described. The tissues are fixed in 10% formalin followed by partial dehydration in 80% iso-propyl alcohol; decalcified in 5% nitric acid in 10% formalin; washed for 5-8 hours in running water and neutralized in 5% aqueous sodium sulfate for 24 hours, followed by further washing in running water for another 24 hours. They are then dehydrated over a period of 4 days in increasing grades of iso-propyl alcohol and embedded in tissue-mat in sub-atmospheric pressure of 10 lb. Sections so obtained were better suited for cytological study than celloidin preparations, and serial sections could be obtained in a much shorter time.     

The effects of clove oil on coral: An experimental evaluation using Pocillopora damicornis (Linnaeus)

Clove oil solution (10% clove oil, 90% ethanol) is an anaesthetic that is widely used to catch demersal fish on coral reefs. This study assessed the effects of clove oil solution on colonies of Pocillopora damicornis, a cosmopolitan reef coral. In the laboratory, low concentrations (0.5 ppt) of clove oil solution had no effect on coral colour or photosynthetic efficiency, irrespective of exposure time (1-60 min). Corals treated with high concentrations (50 ppt) of clove oil solution died immediately, including those that were exposed briefly (1 min). Intermediate concentrations (5 ppt) of clove oil solution produced variable results: a 1 min exposure had no effect, a 10 min exposure caused bleaching and reduced photosynthetic efficiency, and a 60 min exposure caused total mortality. To validate these observations, clove oil solution was applied to corals in situ. Sixty-three days after application, corals treated with 10 ml of clove oil solution appeared to be unaffected. It was concluded that (1) limited amounts of clove oil solution are unlikely to harm this coral, and (2) clove oil solution may represent an ‘eco-friendly’ alternative to cyanide for use in the live reef-fish trade.     

A Controllable Differential Stain for the Hypophysis by Adaptation of the Mallory Triple Stain

A selective and controllable staining method for the hypophysis has been developed with rat material, using Mallory's triple stain as a basis.

Fix in Zenker neutral formol for 6 hours. Longer fixation is undesirable. Transfer to 30% alcohol plus a few drops of a saturated solution of I₂ in aqueous KI over night. Gradually complete dehydration and clear in cedar oil. Infiltrate with a paraffin mixture (paraffin, rubber-paraffin, bayberry wax and beeswax). Section 3-Sμ. Hydrate to distilled water, placing a few drops of a KI-I₂ solution in the 50% alcohol. Stain in 1% acid fuchsia for 30 minutes. Rinse, and differentiate in a weak NH₄OH solution (one drop 28% NH₄OH to 200 cc. HOH). When differentiation is complete, transfer to a 0.5% phosphomolybdic acid solution for 3 minutes, after first stopping the differentiation with a 0.1% HC1 solution and then rinsing with distilled water. Stain for one hour in a solution of: 1% anilin blue, water soluble, 2% orange 6, and 1% phosphomolybdic acid. Rinse in distilled water plus a few cubic centimeters of the stain. Differentiate in 95% alcohol, transfer to absolute alcohol and clear in a mixture of 30% oil of cedar, 40% oil of thyme, 15% absolute alcohol and 15% xylene. Finally, transfer to xylene and mount.     

An Improved Method of Decalcification

Following several experimental investigations, an improved method of decalcification has been devised. The principle of this decalcification method is to obtain complete decalcification by a mixture of as high pH as possible without diminishing the stainability of the Nissl-granules (with Einarson's progressive staining method by means of gallocyanin). This is accomplished by the help of a buffer solution of equal parts of 8 N formic acid and 1 N sodium formate (pH 2.2). After-treatment consists only in rinsing in flowing water for 24 hours. Dehydration is in alcohol (70%, 96%, 100%); cedar oil; ligroin. Embedding in paraffin follows.