Permanent Mounting of Unstained and Aceto-Orcein Stained Cells in the Water-Soluble Medium, Abopon

For cellular morphology, mammalian cells were grown on cover slips in Leigh ton tubes, fixed in 1% osmic acid vapor for 2 min, decolorized with 30% H₂O₂ in 5% ammonium oxalate solution (1:7) for 2 min, then washed thoroughly, and finally mounted in a water-soluble medium consisting of a saturated solution of Abopon in 0.2 M phosphate buffer, pH 7.0. For chromosomal analysis of similarly cultured cells, aceto-orcein preparations were made by conventional methods, with the following minor modifications: following pretreatment with colchicine and hypotonic expansion, the cells on the cover slips were fixed in acetic-alcohol (1:3), air dried, incubated at 37° C for 15 min in 2% orcein in 45% acetic acid, rinsed in 45% acetic acid, washed several times in distilled water, and finally mounted in Abopon mounting medium. Both kinds of preparations were allowed to harden for 24 hr before being handled. Such slides will keep for years at room temperature. Studies requiring frequent comparisons of cellular and chromosomal morphology of cultured cells can thus be extended over long periods of time.     

Permanent Mounting of Unstained and Aceto-Orcein Stained Cells in the Water-Soluble Medium,Abopon

For cellular morphology, mammalian cells were grown on cover slips in Leigh ton tubes, fixed in 1% osmic acid vapor for 2 min, decolorized with 30% H₂O₂ in 5% ammonium oxalate solution (1:7) for 2 min, then washed thoroughly, and finally mounted in a water-soluble medium consisting of a saturated solution of Abopon in 0.2 M phosphate buffer, pH 7.0. For chromosomal analysis of similarly cultured cells, aceto-orcein preparations were made by conventional methods, with the following minor modifications: following pretreatment with colchicine and hypotonic expansion, the cells on the cover slips were fixed in acetic-alcohol (1:3), air dried, incubated at 37° C for 15 min in 2% orcein in 45% acetic acid, rinsed in 45% acetic acid, washed several times in distilled water, and finally mounted in Abopon mounting medium. Both kinds of preparations were allowed to harden for 24 hr before being handled. Such slides will keep for years at room temperature. Studies requiring frequent comparisons of cellular and chromosomal morphology of cultured cells can thus be extended over long periods of time.     

Chromosome Preparatons of Bone Marrow Cells without Prior In Vitro Culture or In Vivo Colchicine Administration

Bone marrow (about 0.5 ml) from au erythropoietic region is freed of blood clots by washing 1-3 min in 1 μg/ml colchicine solution (2-3 ml) and then soaking 1-2 hr at 20-30° C in a second change. For mammalian or avian marrows, the colchicine is made up in phosphate-buffered (pH 7) physiological NaCl solution; for amphibian, Ringer's A solution. Next the specimens are soaked about 20 min in a hypotonic solution as follows: for mammalian, 1% Na-citrate; for avian, a 1:4 dilution of the buffered NaCl solution by distilled water; and for amphibian, Ringer's A-distilled water, 1:1. Then they are heated in a mixture of 2% orcein in 45% acetic acid and 1 N HCl, 9:1. Immediately after heating, squash preparations are made with 2% acetic-orcein in the usual manner. An alternative method is to dissociate the marrow cells by agitating after colchicine treatment. Then, recovering the cells between changes by low-speed centrifugation, to carry out the hypotonic treatment and subsequent fixation in Carnoy's solution I (alcohol acetic, 3:1) before drying the cells onto slides from the fixative. After thorough drying the slides may be stained 10-20 min in acetic orcein, or by other suitable technics.     

A micromethod for chromosome preparation from individual hematopoietic colonies cultured in methylcellulose

We report a micromethod for chromosome preparation from individual hematopoietic colonies cultured in methylcellulose. The entire process was carried out on poly-Lysine (PL)-coated slides. Individual colonies were transferred into 10 microliter of 0.075 M KCl and placed on PL-coated slides. After hypotonic treatment of the colony cells and their attachment to the slides, the cells were fixed by a three-step procedure as follows: addition of a 30% fixative (3:1 methanol:acetic acid) diluted with the hypotonic solution, addition of 20% ethanol, and subsequent immersion of the slides in a 100% fixative. The slides were flame dried and Giemsa stained. Q- and G-banding techniques also were used. These procedures provided analyzable chromosome preparations, even from colonies containing fewer than 50 cells.     

Carbol Fuchsin as A Stain for Human Chromosomes

Cells derived from cultures of bone marrow or leucocytes were treated with hypotonic citrate solution, squashed in 45% acetic acid frozen with CO₂ to allow removal of the cover glass without disturbing the smear, and stained by the following schedule: absolute alcohol, 5 min; coat with 0.2% parlodion and air dry; 70% alcohol, 5 min; distilled water, 5 min; stain 2-5 min in a mixture of 45 ml of a 0.3% solution of basic fuchsin in 5% phenol, 6 ml of glacial acetic acid, and 6 ml of 37% formaldehyde. Differentiate and dehydrate in absolute alcohol, clear in xylene and cover. The stain is durable for several weeks if slides are stored in darkness when not in use. Results resemble those obtained by Feulgen or aceto-orcein methods.     

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).     

Preservation of large-scale chromatin structure in FISH experiments

The nuclear organization of specific endogenous chromatin regions can be investigated only by fluorescence in situ hybridization (FISH). One of the two fixation procedures is typically applied: (1) buffered formaldehyde or (2) hypotonic shock with methanol acetic acid fixation followed by dropping of nuclei on glass slides and air drying. In this study, we compared the effects of these two procedures and some variations on nuclear morphology and on FISH signals. We analyzed mouse erythroleukemia and mouse embryonic stem cells because their clusters of subcentromeric heterochromatin provide an easy means to assess preservation of chromatin. Qualitative and quantitative analyses revealed that formaldehyde fixation provided good preservation of large-scale chromatin structures, while classical methanol acetic acid fixation after hypotonic treatment severely impaired nuclear shape and led to disruption of chromosome territories, heterochromatin structures, and large transgene arrays. Our data show that such preparations do not faithfully reflect in vivo nuclear architecture.     

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.     

An improved method for cytogenetic analysis of meiotic aneuploidy in rodent and frog spermatocytes

Methods are described for the attachment of isolated spermatocytes to glass slides and the subsequent hypotonic swelling and gradual fixation of the metaphase I and metaphase II cells. The methods minimize cell loss and cell disruption and meiotic metaphase chromosomes become spread within residual cytoplasm thus reducing artefactual chromosome loss. Metaphase II complements from mouse, rat and frog spermatocytes prepared by these procedures had relatively low frequencies of hypoploidy (0.5-1.6%). Bivalent loss was not detected in 916 metaphase I complements. Injection of 0.1 mg/kg demecolcine into mice increased the incidence of metaphase II hypoploidy 8-fold. The hypoploid and hyperploid frequencies here increased equally. The results suggest that the methods described may be useful for the analysis of mechanisms of meiotic aneuploidy including aneuploidy resulting from chromosome loss during meiosis I.     

Chromosome Banding Technique with Trypsin-Giemsa in Plants

The present paper is the investigation for the chromosome banding technique of trypsin-Giemsa in plants. Vicia faba, Allium cepa and Secale cereale were used. The fixed roots were hydrolysed in 0.1 N HCl or 45% acetic acid and squashed in a drop of 45% acetic acid. Coverslips were removed by the freezing method and the slides dried in the air. The preparations were then immersed in 0.025% solution of trypsin in phosphate-buffered saline (pH 7.2) at 34 ℃ for 10–15 minutes. Staining was performed on 10% Giemsa at pH 7.2. This method has some unique features. It is fast and technically simple and C-banding of good-quality can be consistently demonstrated. In this paper the chromosome banding pattern with trypsin-Giemsa in Vicia faba is analyzed and the mechanism of chromosome banding with trypsin-Giemsa is also discussed.     

Leukocytes Cultured from Small Inocula of Whole Blood and the Preparation of Metaphase Chromosomes by Treatment with Hypotonic KCL

Leukocytes were cultured from 0.2 ml of whole blood inoculated into 5 ml portions of a medium consisting of Eagle's basal amino acids and vitamins at double strength in Earle's balanced salt solution brought to pH 7.0 with 7.5% NaHCO₃, and containing additives: glutamine, 2 mM; penicillin, 100 units/ml; streptomycin, 100 μg/ml; phenol red, 7 μg/ml; fetal or newborn agammaglobulin bovine serum, 15%; phytohemagglutinin M, 2%; and U.S.P. heparin sodium, 20,000 units/liter. Cultures were incubated in closed 60 × 28 mm screw-cap vials, in a gas phase initially of room air, for 3 days at 37 C, with colchicine to make 0.2 μg/ml added for the final 3-5 hr. After incubation, the cells were separated from the medium by centrifugation, the medium replaced by 0.075 M KCI plus 16 U.S.P. units/ml of heparin sodium at 37 C, cells resuspended and allowed to incubate 10 min. Removal of the hypotonic KCI was followed by fixation in methanol-acetic acid, 3:1 (changed twice), spreading cells on slides by the air-drying method, and staining with 1% natural orcein (G. T. Gurr) in 60% acetic acid. Dehydration and covering completed the preparation. KCI, 0.075 M, has been used advantageously in the above way and for cells cultured by other means from skin and other organs of man and other mammals. Combined advantages of the method are: culture of leukocytes from small volumes of whole blood, with very few failures to obtain mitotic cells; a medium which can be stored frozen in culture vials, and which in a simpler form is usable for long term culture of other cell types; and the use of KCI for hypotonic treatment.     

Novel Fixation of Plant Tissue, Staining through Paraffin with Alcian Blue and Hematoxylin, and Improved Slide Preparation

Onion (Allium cepa) root tips were fixed in a proprietary solution without aldehyde, toxic metals or acetic acid. Fixed specimens were embedded in paraffin, sectioned on a rotary microtome and mounted on detergent-washed slides without adhesive. Slides with ribbon segments affixed were immersed in 0.2% aqueous alcian blue 8GX in screw-capped Coplin jars in a water bath at 50 C for 1 hr. Excess alcian blue was rinsed off under cold running tap water and the slides were immersed in quick-mixed hematoxylin at room temperature for 15 min. Stained slides were deparaffinized, rinsed with isopropanol, air dried, and coverslips were affixed with resin. Thus, the traditional paraffin microtechnique has been modified at all steps from fixation to finishing slides with coverslips.     

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.     

Chromosome preparations in the field from mammals long after death

Chromosome preparations of high quality can be obtained from bone marrow cells of small mammals that have been dead for 20 hr or longer. The bone marrow is rinsed out of the femurs with RPMI medium supplemented with 15% fetal calf serum. Add 0.05-0.1 ml of a 0.01% colchicine solution to 5 ml of medium-cell suspension. After 1/2-1 hr of colchicine treatment at 37 C the cells are spun down and the supernatant replaced by 5 ml of hypotonic (0.075 M) KCl. After 12 min in the hypotonic solution at 37 C the cells are fixed in methanol:acetic acid 3:1. Air dried preparations are made after repeating the fixation procedure three times and the chromosomes are stained with Giemsa, if required after pretreatment of the preparations for banding, e.g., GTG. Technical hints for field work are given. The technique has proven successful with several species of rodents and shrews.     

A Phloxine-Azure-Hematoxylin Sequence for Differential Staining of Cells in Pancreatic Islets

Sections of 6 μ from tissues fixed in Susa or in Bouin's fluid (without acetic acid) and embedded in paraffin were attached to slides with Mayer's albumen, dried at 37 C for 12 hr, deparaffinized and hydrated. The sections fixed in Susa were transferred to a I₂-K₁ solution (1:2:300 ml of water); rinsed in water, decolorized in 5% Na₂S₂O₃; washed in running water, and rinsed in distilled water. Those fixed in Bouin's were transferred to 80% alcohol until decolorized, then rinsed in distilled water. All sections were stained in 1% aqueous phloxine, 10 min; rinsed in distilled water and transferred to 3% aqueous phosphotungstic acid, 1 min; rinsed in distilled water; stained 0.5 min in 0.05 azure II (Merck), washed in water; and finally, nuclear staining in Weigert's hematoxylin for 1 min was followed by a rinse in distilled water, rapid dehydration through alcohols, clearing in xylene and covering in balsam or a synthetic resin. In the completed stain, islet cells appear as follows: A cells, purple; B cells, weakly violet-blue; D cells, light blue with evident granules; exocrine cells, grayish blue with red granules.     

A Naphthol Yellow S and Erythrosin B Staining Procedure for Use in Studies of the Acrosome Reaction of Rabbit Spermatozoa

Rabbit spermatozoa suspended in Krebs-Ringer-phosphate containing 0.25% glucose were smeared on polylysine-coated slides and dried in air at room temperature for 2 hr to overnight. Smears were stained in 0.1% naphthol yellow S in 1.0% acetic acid for 30 min at room temperature, blotted, rinsed in 1.0% aqueous acetic acid for 10-15 sec, drained and stained for 7 min in a mixture of equal parts of aqueous naphthol yellow S and erythrosin B (final concentration of each dye 0.1% w/v) at pH 4.6-5.0 (pH adjusted with acetic acid). Stained slides were well rinsed in distilled water adjusted to pH 4.65.0 with acetic acid, blotted, allowed to dry completely, rinsed in xylene and mounted in synthetic resin. Acrosomal caps were stained cherry-red (apical ridge) to pink (dorsal and ventral aspects); postnuclear caps stained pale pink; nuclei were either unstained or stained a very faint yellowish-pink. The mid-piece and flagellum were stained different shades of pink. The procedure is simple, rapid, and gives highly reproducible results. When present, acrosomes are easily detected regardless of the density of the smear.     

A naphthol yellow S and erythrosin B staining procedure for use in studies of the acrosome reaction of rabbit spermatozoa.

Rabbit spermatozoa suspended in Krebs-Ringer-phosphate containing 0.25% glucose were smeared on polylysine-coated slides and dried in air at room temperature for 30 min at room temperature, blotted, rinsed in 1.0% aqueous acetic acid for 10-15 sec, drained and stained for 7 min in a mixture of equal parts of aqueous naphthol yellow S and erythrosin B (final concentration of each dye 0.1% w/v) at pH 4.6-5.0 (pH adjusted with acetic acid). Stained slides were well rinsed in distilled water adjusted to pH 4.6-5.0 with acetic acid, blotted, allowed to dry completely, rinsed in xylene and mounted in synthetic resin. Acrosomal caps were stained cherry-red (apical ridge) to pink (dorsal and ventral aspects); postnuclear caps stained pale pink; nuclei were either unstained or stained a very faint yellowish-pink. The mid-piece and flagellum were stained different shades of pink. The procedure is simple, rapid, and gives highly reproducible results. When present, acrosomes are easily detected regardless of the density of the smear.     

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.     

Fast direct method of obtaining metaphase and prometaphase chromosomes from chorion biopsy cells and human embryos during the lst semester of pregnancy]

Samples of chorionic villi and embryonic tissues (brain, brain--sheaths) are thoroughly washed with Hank's solution, immediately subjected to hypotonic treatment (0.9% sodium citrate plus few drops of 0.01% colchicine) 37 degrees C, 30 min, prefixed 20 min with equal amount of standard fixative mixture, twice fixed in standard fixative solution (1 hour, -10 degrees C), hydrated with equal volume of distilled water (5-10 min), dried, macerated directly on the slide with 60% acetic acid. The cell suspension is then evenly spread on the slide surface, dried, postfixed and stained. The method provides sufficient amount of metaphase and prometaphase mitotic plates suitable for differentiating staining in 1.5-2 hours after sampling and might be recommended for routine chromosomal analysis in prenatal diagnosis of inherited diseases during early pregnancy.     

Stains for Strands in Sieve Tubes

Two very simple procedures give a staining-fixation of the so-called “strands” as well as portions of the sieve plates of sieve tubes of various broad-leaved deciduous trees. One procedure consists of placing hand-made sections (radial or tangential) of inner bark for 5 min in a 0.2% solution of ponceau S in 3% trichloroacetic acid, then soaking 5 min in 5% acetic acid. A second procedure consists of placing sections in 0.001% nigrosin in 2% acetic acid for approximately 15 hr, then washing briefly in distilled water. In the former procedure, strands, sieve plates, and what appears to be plasmalemma, appear reddish or pink, while cell walls do not stain. In the latter, strands and sieve plates appear bluish but phloem cell walls also become bluish, although xylem cell walls usually remain unstained.