Embedding Media for 1-2 Micron Sectioning. 2. Hydroxyethyl Methacrylate Combined with 2-Butoxyethanol

A hydroxyethyl methacrylate monomer medium incorporating 2-butoxyethanol requires 2 stock solutions for embedding. Solution A: 80 ml of hydroxyethyl methacrylate (Rohm and Haas Co., Philadelphia, Pa.) is mixed well with 16 ml of 2-butoxyethanol; 0.27 gm of benzoyl peroxide, the catalyst, is added and permitted to dissolve. Heating to 40-50 C may be used to accelerate its solution. Solution B: polyethylene glycol 200 or 400, 15 parts, and N,N-dimethylaniline, 1 part, are mixed thoroughly. Tissues are dehydrated in the customary manner to absolute ethanol or other comparable dehydrant, infiltrated completely with A, then cast in a mixture consisting of 42 parts of A well mixed with 1 part of B. Polymerizaion occurs in 4-7 hr. In a water bath at 20 C the time required was about 7 hr; at 28 C, 4 hr. This medium is based on the author's water-polyethylene glycol-hydroxyethyl methacrylate monomer medium (Stain Techn., 42: 119-23, 1967).     

Hydroxyethyl Methacrylate Combined with Polyethylene Glycol 400 and Water; an Embedding Medium for Routine 1-2 Micron Sectioning

Pieces of tissue, with the largest dimension not exceeding 7 mm, are fixed and dehydrated by the procedures of choice. Two stock solutions: A, for infiltration; and B, the accelerator, are used in embedding. Formulas: A, 80 ml of glycol methacrylate (2-hydroxyethyl methacrylate—Rohm and Haas Co., Philadelphia, Pa.) is mixed well with 12 ml of polyethylene glycol (Carbowax) 400 and 8 ml of water; then 0.27 gm of benzoyl peroxide added, heated to dissolve the peroxide, and allowed to cool to room temperature. B, polyethylene 200 or 400, 15 parts, and N,N-dimethylaniline, 1 part, mixed thoroughly. Tissues are first infiltrated completely with solution A, then cast in a mixture consisting of 42 parts of A mixed with 1 part of B. Polymerization occurs in 45 min to 3 hr, depending on the temperature. In a water bath at 20 C, the time required was found to be about 3 hr; at 25 C, 1.5 hr; and at 30 C, 45 min. The plastic block can be trimmed easily, and sections 1-2 μ thick readily cut. Sections can be attached to slides by water flotation, without adhesive, and should be dried at room temperature. Staining with aqueous solutions of basic and acid dyes, without removing the embedding matrix, is sharp and brilliant. When staining of the matrix by basic dyes occurs, this background stain can be completely removed by differentiating in either 2-butoxyethanol, pure ethanol, or a mixture of the two. A number of histochemical reagents have been found compatible with this embedding procedure.     

Serial Sections of Cooked Rice Embedded in Carbowax

Serial sections of cooked rice kernels may be obtained by following either of two dehydration schedules and embedding in Carbowax. In the first schedule the cooked, rinsed and drained kernels are immersed several days in a nonaqueous fixative composed of: isopropyl alcohol, 10 ml; propionic acid, 30 ml; acetone, 10 ml; methylal, 40 ml; dioxane, 30 ml; and propylene glycol, 30 ml (Newcomer's, modified), followed by 7 or 8 days in equal parts of propylene glycol, dioxane and glycerol (changed once), and 4 days on a warming table in the same mixture with 5% Carbowax added. The dehydrated kernels are then infiltrated 4-24 hr with a Carbowax embedding mixture. In the second schedule they are immersed several days in an aqueous solution consisting of: propylene glycol, 12.5 ml; polyethylene glycol 400, 12.5 ml; either with 75 ml of water containing 0.1% thymol, or with a mixture of water, 65 ml; formalin, 10 ml; CaCl₂, 1 gm; and CdCl₂, 1 gm; followed by 3 or 4 days in 50% propylene glycol, and 3 or 4 days on a warming table in 80% propylene glycol with 5% Carbowax added. Infiltration is as above. The composition of the embedding mixture is varied according to the temperature and humidity likely to prevail during sectioning. The texture of the wax may be improved by adding small amounts of gum arabic, spermaceti, and glycerol. Serial sections 3-10 μ thick are placed on clean dry slides, and adhesive dropped at the edges of the ribbon of Carbowax until it is dissolved. The adhesive consists of water-glass (concentrated solution), 1 ml; concentrated ammonia, 1 ml; Carbowax, 5 gm; and water, 98 ml. After the slides are dry they are stored, or immersed 10 min in chloroform, collodionized, and passed to staining solutions. Atmospheric conditions affect not only the Carbowax, but also the response to reagents of cooked rice and of sections.     

Leukocyte Culture and Chromosome Preparations from Adult Dog Blood

Plasma is obtained from dog blood after 3 hr settling in a syringe. Portions of the plasma (0.5-1.0 ml) are added to 4 ml of a medium consisting of 17 parts of BME Spinner, 3 parts of calf serum, 0.5 parts of glutamine, 0.5 parts of penicillin-streptomycin, and 0.1-1.0 parts of Scarlet Runner bean phytohemagglutinin. Colchicine, 0.1 ml of 10:1 stock solution, is added after 72 hr and incubation continued for 2 hr, then centrifuged 5 min at 700 rev/min. The supernatant is discarded, 3 ml of distilled water added, and the cell suspension centrifuged again. The supernatant is discarded and the fixative, consisting of 45% glacial acetic acid allowed to act for 0.5 hr. Acetic-orcein stains of smears were very satisfactory.     

A Plastic Embedding Medium for Thin Sectioning in Light Microscopy

Tissue blocks with surface areas up to 2 cm² can be sectioned at 1 or 2 μ after embedding in a medium consisting of: methyl methacrylate, 27 ml; polyethylene glycol distearate MW 1540, 6 gm; dibutyl phthalate, 4 ml; and Plexiglas molding powder A-100, 9 gm (added last). The methacrylate mixture is polymerized at 50° C by benzoyl peroxide, 0.8 gm/ 100 ml of methacrylate. The polymerized matrix is transparent and the blocks can be cut on a rotary microtome with a steel knife. The plastic can be removed from sections with acetone prior to staining. Artifacts caused by embedding and sectioning are negligible     

Quantitative autoradiography of 3H-thymidine-labelled nuclei: Reduction of grain count caused by washing in water

Summary Ehrlich ascites tumour cells and epithelial cells of intestinal crypts were labelled in vitro and in vivo with the pulse of either tritium or carbon-14-thymidine. Fixed cells were washed in either distilled or tap running water for 1/6–24 hr and autoradiographed. Quantitation of autoradiographs showed that the washing of cells labelled with tritiated thymidine caused remarkable diminution of grain count. The cells labelled with carbon-14-thymidine were not affected by washing. The strict control of washing in water of tritium-labelled specimens is recommended.     

Silver Impregnation of Degenerating Axons in the Central Nervous System: A Modified Technic

Frozen sections of formalin-fixed brains containing surgical lesions, were treated with 15% ethanol for 0.5 hr., soaked in 0.5% phosphomolybdic acid for 0.25-1.0 hr., and subsequently treated with 0.05% potassium permanganate for 4-10 min. (The duration of the latter treatment is critical and individually variable). Subsequent procedure is as follows: decolorize in a mixture of equal parts of 1% hydroquinone and 1% oxalic acid; wash thoroughly and soak sections in 1.5% silver nitrate for 20-30 min.; ammoniacal silver nitrate (silver nitrate 0.9 g., distilled water 20 ml., pure ethanol 10 ml., strong ammonia 1.8 ml., 2.5% sodium hydroxide 1.5 ml.) 0.5-1.0 min.; reduce in acidified formalin (distilled water 400 ml., pure ethanol 45 ml., 1% citric acid 13.5 ml., 10% formalin 13.5 ml.) 1 min.; wash, and pass section through 1 % sodium thiosulf ate (0.5-1.0 min.); wash thoroughly and pass sections through graded alcohols and xylene (3 changes); cover in neutral synthetic resin.     

Histochemical Demonstration of Succinic Dehydrogenase in Ascites Tumor Cells

Fresh undiluted tumor ascites (0.05 ml) withdrawn from peritoneal cavity was placed immediately in a centrifuge tube containing 2.0 ml of an aqueous mixture prepared with 1 part each of the following solutions: 1% neotetrazolium chloride, 0.2 M sodium succinate and 0.1 M phosphate buffer, pH 7.4. The tube was incubated for 2 hr at 37°C and centrifuged for 3 min at 700 rev/min. The precipitate was washed with 0.85% saline solution and subsequently fixed with neutral 10% formalin for 10 min. After centrifugation, smears or squash preparations of the precipitate were prepared. Succinic dehydrogenase activity was demonstrated very distinctly and uniformly by the granular deposition of a deep purple pigment intracellularly.     

A Pollen Grain Squash Technique for Saccharum and Related Genera

Anthers collected between 9 and 10 AM were treated for 1 hr at 26-28 C with a 0.5% solution of colchicine, washed for 2-4 min in water, placed in 0.002 M 8-hydroxyquinoline for 1 hr, washed in water for 10 min and fixed in: methanol, 60 ml; chloroform, 30 ml; distilled water, 20 ml; picric acid, 1 gm and mercuric chloride 1 gm, for 24 hr. After washing they were hydrolysed in 1 N HCl for 15 min at 60 C, stained in leuco basic fuchsin for 30 min, then smeared on a slide in a drop of acetocarmine. The slides were sealed, stored overnight, the paraffin was removed, and the slide passed through a 1:1 mixture of n-butyl alcohol and acetic acid, then through pure n-butyl alcohol and mounted in Canada balsam. The significant features of this procedure are: (1) use of chromosomes in the haploid condition for karyotype analysis, (2) better exaggeration of constrictions for easier interpretation of chromosome types and (3) good spreading in plants with a large chromosome number.     

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).     

India Ink-Gelatin Vascular Injection of Skeletal Tissues

Dogs under sodium pentothal anesthesia were given 100 mg of heparin intravenously and, 15 min later, exsanguinated. A mixture consisting of Higgins India ink #4415, 1 part, and a 14% solution of calfskin gelatin, 1 part, was injected (at 37° C) into the femoral artery of a disarticulated hindlimb. A pump-type 5 ml hypodermic syringe was connected with a gravity-feed reservoir that contained the injection mass, with a filter in the outflow from the reservoir. Veins severed by the disarticulation of the limb permitted a free flow of the injection fluid out of the vascular tree. After injection of about 400 ml of fluid by pumping the syringe, the limb was chilled (—20° C) to solidify the gelatin. Specimens from cleaned bones were sawed out and fixed in Zenker-formalin solution for 24 hr, then decalcified with 6% sulfosalicylic acid. The technique was completed by embedding in celloidin and sectioning. Blood vessels in both cortical and trabecular regions were completely filled by the injection mass.     

Simplified Culture and Chromosome Preparations of Primate Leukocytes

Plasma recovered from 1 ml of primate peripheral blood by centrifugation is planted in a medium consisting of 80% TC-199 and 20% fetal bovine serum, to which 0.125 ml of phytohaemagglutinin/5 ml is added. The pH is adjusted to 7 with 10% NaHCO₂. The mixture is incubated 68 hr, Colcemide to give 1 μg/ml is added, and incubation continued for 4 hr. Following centrifugal separation, the cells are given a hypotonic treatment with 0.75% sodium citrate for 15 min, then centrifuged again and fixed in 3:1 methanol-glacial acetic acid, 3 changes. Tiny drops of the cell suspension are placed on a slide, spread by blowing, and air dried. The preparations are stained with 15% Giemsa solution in methyl alcohol. The method has been successfully used in 256 specimens from 25 different species.     

Leucocyte Culture and Chromosome Preparations from Pig Blood

Blood was drawn into heparinized tubes from any large vein and allowed to settle 2-3 hr at 3-5 C. The cell sample consisting of 1 ml drawn from the buffy coat and 2 ml from the plasma was planted in the following medium: Medium 199 (Difco), 10 ml; penicillin G sodium, 1000 USP units; dihydrostreptomycin, 1 mg; and Bacto-PHA-M (Difco), 0.2 ml. Incubation, with twice daily shaking, was at 37 C for 68-70 hr; colchicine to give 4 μ ml was then added and incubation continued for 3-4 hr. The bulk of the medium was removed by centrifugation, the cells washed once in Hanks' salt solution, centrifuged, and all but 0.5 ml of the fluid decanted; 1.5 ml of distilled water at 37 C was added, the cell suspension incubated at 37 C 5-15 min, followed by centrifugation and fixation in methanolacetic acid 3:1 (3 changes) as usual. Spreads were made by applying 4-5 drops of cell suspension to ice-cold slides and burning off the fixative. Giemsa stain was used. The method has proved very satisfactory for determining chromosome numbers in the domestic pig. This number, as determined in 690 cells from Poland China and Duroc gilts and crosses of these breeds was 38 in 611 (88.6%) of the cells.     

Immunofluorescence Microscopy of Cyanurated Tissues

Test tissues consisted of: (1) popliteal lymph nodes of rabbits, removed 6 hr after injection of hind footpads with 0.2 ml of 125 mg/ml solution of 5× crystallized chicken ovalbumin, and (2) lungs from guinea pigs, passively sensitized with rabbit antiovalbumin serum, then anaphylactically shocked by intracardial injection of a 1% chicken ovalbumin solution. Similar control tissues from normal rabbits, and lungs of passively sensitized guinea pigs, but shocked with histamine instead of ovalbumin, were included. Pieces of fresh tissue not exceeding 2 mm³ were fixed as follows: (1) Cyanuration—lymph nodes, 1 hr; lung, 0.5 hr; both at 23-27 C—in anhydrous methanol containing 0.5% w/v cyanuric chloride and 1% v/v N, N-diethylaminoethanol. (2) Control fixatives—all specimens 18-24 hr at 4—6 C—absolute methanol; 95% ethanol; neutral buffered 10% formalin; and an FAA mixture (formalin, conc., 6; glacial acetic acid, 2; 30% ethanol, 92). Freeze-dried material was either left unfixed (a control) or fixed in xylene or toluene containing 0.5% w/'v cyanuric chloride and 1% v/v N, N-diisopropylaminoethanol; time and temperature as for fresh tissues. All tissues were routinely dehydrated, cleared, and vacuum embedded in an ester wax, diethylene glycol distearate, or in paraffin at 52 C. Sections 2-4 μ thick were attached to gelatin-coated slides, the wax removed in petroleum ether, and stained 20 min at 23-27 C in a 0.10% solution of fluorescein isothiocyanate-conjugated rabbit antiovalbumin globulin, washed in phosphate buffered saline 10 min, dehydrated, cleared and covered in a nonfluorescent medium. With ultraviolet illumination, brightly immunofluorescent, anti-genically specific staining was obtained in cyanurated fresh and freeze-dried lymph node and lung tissues. In contrast, specific staining was diminished or absent in comparable tissues reacted in the control fixatives.     

Staining Reticulin with Gold

This bromine-iodine-gold chloride-reduction sequence stains reticulin in formalin-fixed paraffin sections without risk of sections becoming detached. After hydration, sections are exposed to 0.2% bromine water containing 0.01% KBr for 1 hr, then rinsed and placed for 5 min in a solution consisting of KI, 2 gm; iodine crystals, 1 gm; and distilled water, 100 ml. After this the sections are well washed in distilled water, immersed for 5 min in 1% w/v aqueous solution of chloro-auric acid, again rinsed in distilled water, and the gold is reduced by placing in freshly made 3% H₂O₂ for 2-4 hr at 37 C, or in 2% oxalic acid for 1-3 hr at the same temperature.     

A Modified Pyridine-Silver Stain for Teased Preparations of Motor and Sensory Nerve Endings in Skeletal Muscle

This technique has been developed especially to stain sensory receptors which have been localised intramuscularly by electrophysiological means. Rat intertransverse caudal muscles, removed immediately after death, are fixed for 24 hr in a freshly prepared mixture of absolute ethyl alcohol, 4.5 ml; distilled water, 5 ml; and concentrated HNO_a, 0.1 ml. After a further 24 hr in 10 ml of absolute ethyl alcohol containing 0.1 ml of ammonia solution (sp. gr. 0.88), the muscles are washed in distilled water for 30 min and placed in full strength pyridine for 2 days. They are then washed for 24 hr in distilled water (changed 5-8 times) and left in 2% AgNO₃, in the dark for 3 days at 25 C. Following reduction in 10 ml of 5% formic acid containing 0.4 gm of pyrogallol for 6-24 hr, the specimens are washed briefly in distilled water and stored in pure glycerol. The nerve endings can then be teased out and mounted in glycerol, under cover glasses ringed with a waterproof cement. The advantage of this method is that it gives consistently good staining of receptors and motor end-plates in small muscles of the rat     

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.     

A Method for Staining Pollen Tubes in Pistil

A quadruple staining procedure has been developed for staining pollen tubes in pistil. The staining mixture is made by adding the following in the order given: lactic acid, 80 ml; 1% aqueous malachite green, 4 ml; 1% aqueous acid fuchsia, 6 ml; 1% aqueous aniline blue, 4 ml; 1 % orange G in 50% alcohol, 2 ml; and chloral hydrate, 5 g. Pistils are fixed for 6 hr in modified Carnoy's fluid (absolute alcohol:chloroform:glacial acetic acid 6:4:1), hydrated in descending alcohols, transferred to stain and held there for 24 hr at 45±2 C They were then transferred to a clearing and softening fluid containing 78 ml lactic acid, 10 g phenol, 10 g chloral hydrate and 2 ml 1% orange G. The pistils were held there for 24 hr at 45±2 C, hydrolyzed in the clearing and softening fluid at 58±1 C for SO min, then stored in lactic acid for later use or immediately mounted in a drop of medium containing equal parts of lactic acid and glycerol for examination. Pollen tubes are stained dark blue to bluish red and stylar tissue light green to light greenish blue. This stain permits pollen tubes to be traced even up to their entry into the micropyle.     

A method for staining pollen tubes in pistil

A quadruple staining procedure has been developed for staining pollen tubes in pistil. The staining mixture is made by adding the following in the order given: lactic acid, 80 ml; 1% aqueous malachite green, 4 ml; 1% aqueous acid fuchsin, 6 ml; 1% aqueous aniline blue, 4 ml; 1% orange G in 50% alcohol, 2 ml; and chloral hydrate, 5 g. Pistils are fixed for 6 hr in modified Carnoy's fluid (absolute alcohol:chloroform:glacial acetic acid 6:4:1), hydrated in descending alcohols, transferred to stain and held there for 24 hr at 45 +/- 2 C. They were then transferred to a clearing and softening fluid containing 78 ml lactic acid, 10 g phenol, 10 g chloral hydrate and 2 ml 1% orange G. The pistils were held there for 24 hr at 45 +/- 2 C, hydrolyzed in the clearing and softening fluid at 58 +/- 1 C for 30 min, then stored in lactic acid for later use or immediately mounted in a drop of medium containing equal parts of lactic acid and glycerol for examination. Pollen tubes are stained dark blue to bluish red and stylar tissue light green to light greenish blue. This stain permits pollen tubes to be traced even up to their entry into the micropyle.     

Effects of bovine serum proteins in culture medium on post-warming survival of bovine blastocysts developed in vitro

Experiments were conducted to investigate the factors affecting the survival of bovine blastocysts produced in vitro after cryopreservation by vitrification. Zygotes were obtained by in vitro maturation and fertilization of oocytes. Embryos used in this study were developed in vitro at Day 7 and 8 (Day 0 = insemination day) in modified synthetic oviduct fluid medium supplemented with calf serum or BSA. Embryos were cryopreserved in a two-step protocol consisting of exposure to 10% ethylene glycol for 5 min, followed by the original vitrification solution (designated as VS) consisting of 40% (v/v) ethylene glycol, 6% (w/v) polyethylene glycol and 0.5 M sucrose in phosphate-buffered saline for 1 min. After warming, embryos were cultured in modified TCM-199 for an in vitro survival assay. The highest survival rate was obtained from the warmed embryos developed at Day 7 in medium supplemented with BSA (82.6%), and there were significant differences between results with calf scrum and BSA treatment (42.4 and 70.7%, respectively; P < 0.01). However, there were no significant differences in the cell numbers of embryos among the treatments. These results suggest that the survival of embryos developed in medium with BSA is superior to that of embryos developed in medium containing calf serum, although the cell numbers of the embryos developed under both media were similar.