Chromosome Preparations from Mouse Embryos During Early Organogenesis: Dissociation After Fixation, Followed by Air Drying

Embryos are put into 1% sodium citrate at 37 C; 7- and 8-day specimens requiring about 20 min. With increasing age, the duration of treatment is increased up to 50 min. Handling is facilitated by keeping specimens in a small glass vessel for observation under a binocular microscope, and by changing fluids with a fine-tipped pipette. Fixation in ethanol-acetic acid 3:l for 2-3 hr is uncritical, as material may be stored in the fixative overnight at 4 C. Staining in toto with 2% orcein in 50% acetic acid follows, requiring 0.5-1 hr (storage in this solution up to 2 wk at 4 C is permssible). After staining, specimens are subjected to cellular dissociation in a mixture of glacial acetic and 50% lactic acid, the action of which is controlled by the duration of treatment and by increasing the ratio of lactic to acetic from 1:Z (younger embryos) to 3:2 (older embryos). Only 1-3 drops of the dissociating fluid is used for each embryo, to favor concentration of the free-floating cells. Since the time required varies from several minutes to nearly an hour, the most favorable degree of dissociation can best be judged by the cloudiness produced in the dissociating fluid. A small drop not exceeding 2 mm in diameter, of the cell suspension, is placed on a slide and followed immediately by a normal-sized drop of fresh 3:1 ethanol-acetic. After drying, the chromosomes are stained with lactic-acetic-orcein or other suitable stain. The method gives satisfactory results with embryos from the 7th to 11th day of pregnancy.     

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.     

Staining Germinating Pollen and Pollen Tubes

Germinating pollen on stigmas and pollen tubes in styles of Antirrhinum, Brassica, Oenothera, Raphanus, Rosa, solatium and Tagetes spp. were prepared for examination as follows: The styles were fixed in ethyl alcohol-acetic acid 3:1 for 1 hr, and hydrolyzed at 60°C for 5 to 60 min (depending on the species) in 45% acetic acid. The stigma with its attached strand(s) of stigmatoid tissue was then dissected out under a stereoscopic microscope, placed in a few drops of a staining solution made by dissolving 150 mg of safranin O and 20 mg of aniline blue in 25 ml of hot 45% acetic acid. After 5-15 min in this stain, the tissue was placed in a fresh drop of stain on a microscope slide and gently squashed under a cover glass. Because of a gradual precipitation of the aniline blue component, the stain had to be filtered regularly before use. However, a staining solution could be kept at room temperature for several weeks.     

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.     

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.     

Chromosomes of Sequoia sempervirens; 8-Hydroxy-Quinoline-Castor Oil Pretreatment for Improving Preparation

Living root tips of coast redwood were pretreated for 6 hr at 10-14 C in a homogenized mixture of 0.003 M aqueous solution of about 5 ml of 8-hydroxyquinoline and a drop of castor oil. They were rinsed 2-3 times in Carnoy's alcohol-chloroform-acetic acid (3:2:1), and fixation allowed to continue in this fluid for 1 hr at 60 C. They were then hydrolysed in 1 N HC1 at 60 C for 45 min; washed with distilled water, and squashed in a drop of aceto-carmine or aceto-orcein to make a temporary slide, subsequently made permanent by a quick-freezing method. Our work to date seems to confirm 2n = 66 for S. sempervirens.     

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 Squash Technique for Human Male Meiotic Chromosomes: Softening and Concentration of Cells; Mounting in Hoyer's Medium

Human testicular material obtained from autopsies was processed as follows: (1) swelled with 0.3% sodium citrate 1-6 hr; (2) softened with 3 M glucono-delta-lactone 2 hr; (3) stained with aceto- or propiono-carmine overnight; (4) washed in 70% alcohol; (5) macerated in 1:1 alcohol-acetic acid; (6) filtered through gauze to remove tubules and connective tissue; (7) filtrate centrifuged to separate spermatocytes from debris; (8) a drop of supernate from just above the precipitate, containing stained cells, mixed with Hoyer's medium; (9) cover slip applied, preparation warmed, and (10) squashed. Chromosomes remained in good condition for 8 mo.     

Glutaraldehyde-Ammoniacal Silver Carbonate-Formaldehyde Staining of Histones of Mitotic Chromosomes

Cells from monolayer culture of Chinese hamster line Don were treated by Colcemid (0.1 μg/ml) for 2 hr, trypsinized and spun; resuspended in 0.5% sodium citrate solution for 10 min, respun, and then resuspended in a small volume of the supernatant. Slide preparations were made by smearing, followed by air drying for 1 min at room temperature. They were fixed and stained by the following sequence: 2.5% glutaraldehyde in Millonig's buffer, 30 min; distilled water, 6 min, 5 changes; ammoniacal silver at 18-26 C, 10 sec; distilled water, 30 min, 5 changes; 2.5% formalin, 2 min; and distilled water, 3 changes during 15 min. Staining solution: add 225 ml of 5% Na₂CO₃ to 75 ml of 10% AgNO₃, then add concentrated NH₄OH slowly, drop by drop, until the solution is transparent. Finally add 300 ml of dstilled water. Cells treated with cold 0.25 N HCl before fixation were not stained. Sequence modifications show that chromatin does not reduce silver by itself. This method stains the sites of high histone concentrations in mitotic chromosomes of cytogenetic preparations.     

Cytochemical Staining of Sections from Plastic-Embedded Flagellates

Dinoflagellate chromosomes in sections of plastic-embedded cells were stained without removing the plastic. Azur B and Feulgen procedures were used to localise DNA. Azur B was used with Araldite or methacrylate sections by staining in 0.2% stain in 0.05 M citrate buffer at pH 4 for 1 hr at 50 C followed by rinsing in tertiary butyl alcohol to differentiate the chromosomes. Feulgen stain was used with Araldite sections by hydrolyzing in 1 N HCl at 60 C for 10 min, rinsing in water, staining for 24 hr, washing well, drying and covering. Fast green was used with methacrylate sections to stain proteins by flooding the slide with a 0.1% solution of stain in 0.06 M phosphate buffer at pH 8, allowing the stain to dry out at 40-50 C, washing well, drying and covering. Controls were carried out on material fixed in formalin and treated with nucleases or proteolytic enzymes prior to embedding, and staining.     

Counterstaining Golgi-Cox Impregnations with Luxol Fast Blue as a Myelin Stain

Immerse pieces of brain tissue 4 wk in solutions A and B, mixed just before use: A. K₂Cr₂O₇, 1 gm; HgCl₂, 1 gm; boiling distilled water, 85 ml. Boil A for 15 min, cool to 2 C and add: B. K₂CrO₄, 0.8 gm; Na₂WO₄, 0.5 gm; distilled water, 20 ml. Rinse in water and immerse 24 hr in LiOH, 0.5 gm; KNO₃, 15 gm; distilled water, 100 ml. Wash 24 hr in several changes of 0.2% acetic acid and then for 2 hr in tap water. Dehydrate and embed in celloidin. Process a 60 μ section through 70 and 95% ethanol, a 3:1 mixture of absolute ethanol and chloroform, and toluene. Immerse it for 5 min in a solution containing methyl benzoate, 25 ml; benzyl alcohol, 100 ml; chloroform, 75 ml. Orient the section on a chemically clean slide and let air-dry 5-10 min. Process through toluene, 3:1 ethanol-chloroform and 95% ethanol. Place the section for 5-60 min at 60 C in a solution made up of: Luxol fast blue G (Matheson, Coleman and Bell), 1 gm; 95% ethanol, 1000 ml; 10% acetic acid, 5 ml. Hydrate to water and immerse in 0.05% Li₂CO₃ for 3-4 min. Differentiate in 70% ethanol and place in water. Immerse for 5-15 min in a mixture of two solutions: A. cresylechtviolet (Otto C. Watzka, Montreal), 2 gm; 1 M acetic acid, 185 ml; B. 1 M sodium acetate, 15 ml; distilled water, 400 ml; absolute ethanol, 200 ml. Dehydrate to 3:1 ethanol-chloroform. Clear in toluene and apply a coverslip. The technique produces fast Golgi-Cox impregnated neurons against a background of counterstained myelinated fibers. Patterns of the myelinated fibers can be used to localize impregnated neurons.     

Chromosome Spreading Induced by Vegetable Oils

Seeds soaked in the oil extracted from castor beans (Ricinus communis) for 2 hr were germinated in petri dishes on moist filter papers. Root tips were fixed in acetic alcohol (1:3) at 10-14°C, for 24 hr, washed successively with 70% alcohol (15 min) and water (10 min), hydrolysed in 1 N HCl at 60°C for 15 min and stained in leucobasic fuchsin for 30 min. The stained tip was squashed under a cover glass in a drop of acetocarmine and sealed with paraffin wax. The slides were made permanent by separating the cover glass in a mixture of acetic acid and n-butyl alcohol (1:1), passing through 2 changes of n-butyl alcohol and mounting in balsam. Such a method leads to contraction and spreading of chromosomes, without affecting either the clarity of the constriction regions or the anaphase separation of chromosomes.     

A Method for Making Aceto-Carmin Smears Permanent

Anthers are collected and placed in a solution of 1 part acetic acid to 3 parts of absolute alcohol. The contents of the anther are squeezed out on a slide in a drop of Belling's iron-aceto-carmin solution and a cover glass placed over the drop. Care should be taken to remove all anther walls and flower parts. Heat the slide over an alcohol flame for a second, repeating 4 or 5 times. Place the slide in a petri dish filled with a 10% solution of acetic acid. When the cover glass has risen away from the slide gently remove the cover glass and place in a Coplin jar containing equal parts of alcohol and acetic acid. Likewise, place the slide in this solution. Run both cover and slide thru the following solutions: 1 part acetic acid to 3 parts absolute alcohol, 1 part acetic acid to 9 parts absolute alcohol, absolute alcohol and finally equal parts of absolute alcohol and xylol. Recombine the cover and slide in xylol-balsam directly from this solution.     

Handling Germinated Pollen on Millipore Membrane During the Feulgen and Autoradiographic Procedures

To prevent loss of pollen during the Feulgen's procedure, the pollen was grown on an autoclaved membrane filter (Millipore AA WP 025 00) in contact with a sterilized medium containing agar 0.5-1%, sucrose according to the genus (Malus 0.3-0.5 M; Persica and Tulipa 0.4 M), and H₃BO₃, 0.01%. To fix the germinated pollen of most species, the membrane was placed for 2 hr to overnight at 2-4 C on filter paper wet with the following mixture: OsO₄, 1 gm; CrO₃, 1.66 gm; and distilled water, 233 ml. To fix Persica pollen, 10% of glacial acetic acid had to be added to the fixative. Washing with distilled water and bleaching with a mixture of 3% H₂O₂ and sat. aq. ammonium oxalate, 1:1, were performed also on filter paper. Similarly, the preparation was processed for Feulgen staining by use of pieces of filter paper wet with the required fluids. Hydrolysis preceding the Schiff's reagent was performed at room temperature with 5 N HCl for 18 min. The differentiation after the Schiff's action was with 2% K₂S₂O₅ buffered to pH 2.3 with 9 ml of phosphate buffer (KH₂PO₄, 1.4 gm; conc. HCl, 0.35 ml and distilled water to make 100 ml). The stained pollen was floated off the membrane with a drop of glacial acetic acid to a gelatinized or an albumenized slide, and squashed. When the coverslip is removed the preparation may be either dehydrated and mounted or coated with autoradiographic film.     

Whole Mount Preparation of Chick Blastodermal Edge Tissue for Microscopic Examination

Three methods of preparation of chick blastodermal edge tissue for conventional microscopy were attempted: (1) Sections were cut from Water Wax (E. Gurr) embedded material. This method was unsatisfactory due to loss of morphological relationships produced by the inability to attach the tissue to a slide. (2) Frozen sections were cut from embryonic marginal tissue with its underlying white and yellow yolk which was embedded in a 25% gelatin solution. This method retained morphological relationships prior to 24 hr of egg incubation, but was technically impractical in excess of this period of incubation. Also, the gelatin could not be removed from the sections and cellular detail was obscured by its subsequent staining. (3) The blastoderm was removed from the yolk and adherent vitelline membrane, and the yolk dissected from a small piece of this blastodermal edge tissue under a dissecting microscope. The dissected tissue, primarily monocellular and dicellular in thickness, was transferred to a slide and attached to it by allowing a few drops of ether-alcohol (1:1) to flow over it. The plane of the tissue was, therefore, parallel with the plane of the slide. Most fixatives and staining techniques could subsequently be used. Fixing for 30-120 min in ether-alcohol (1:1) containing 0.5% glacial acetic acid gave excellent results with the staining techniques attempted. Fixation with Zenker's fluid for 24 hr followed by a 6 min hydrolysis with 1 N HC1 was best prior to the Feulgen technique.     

A Method for Making Aceto-Carmin Smears Permanent

Anthers are collected and placed in a solution of 1 part acetic acid to 3 parts of absolute alcohol. The contents of the anther are squeezed out on a slide in a drop of Belling's iron-aceto-carmin solution and a cover glass placed over the drop. Care should be taken to remove all anther walls and flower parts. Heat the slide over an alcohol flame for a second, repeating 4 or 5 times. Place the slide in a petri dish filled with a 10% solution of acetic acid. When the cover glass has risen away from the slide gently remove the cover glass and place in a Coplin jar containing equal parts of alcohol and acetic acid. Likewise, place the slide in this solution. Run both cover and slide thru the following solutions: 1 part acetic acid to 3 parts absolute alcohol, 1 part acetic acid to 9 parts absolute alcohol, absolute alcohol and finally equal parts of absolute alcohol and xylol. Recombine the cover and slide in xylol-balsam directly from this solution.     

Prefixing with Paradichlorobenzene to Facilitate Chromosome Study

A technic was developed which resulted in preparations containing many mitotic divisions with chromosomes well fixed and stained, rod-shaped, and spread throughout the cell. This technic has given good results with guayule (Parthenium argentatum), Crepis, Allium, Pisum, Lycopersicon, Tradescantia, and other plants. Material is prefixed in a saturated solution of paradichlorobenzene for 1-4 hours, fixed in 65% acetic acid (or other suitable fixative) for 12-24 hours, hydrolyzed in 10% HCl for 10-30 minutes at 60° C, rinsed in water, transferred to a drop of 45% acetic acid on a slide, and smeared and stained in aceto-orcein. The preparation may be made permanent by separating slide and cover glass in 1 part glacial acetic acid to 1 part absolute alcohol, putting them in absolute alcohol, and then recombining them with a drop of euparol.     

Selection for Electron Microscopy of Specific Areas in Large Epoxy Tissue Sections

Tissue blocks 2 × 2 × 0.4 cm were fixed 6-24 hr in phosphate-buffered 6% glutaraldehyde then sliced to 2 × 2 × 0.1 cm and rinsed in phosphate buffer for at least 12 hr. Fixation was continued for 2 hr in phosphate-buffered 1-2% OsO₄. The slices were dehydrated, infiltrated with Araldite, and embedded in flat-bottomed plastic molds. Sectioning at 4-8 μ with a sliding microtome was facilitated by addition of 10% dibutylphalate to the standard epoxy mixture. The sections were spread on water and attached to coverslips by drying, then heating to 80 C for 1 min. Staining 2 min with 1-3% KMnO₄ and temporary mounting in glycerol on a slide allowed the desired area for electron microscopy to be selected and marked. This area was then cemented to the facet of a conventional epoxy casting with a drop of epoxy resin (without added dibutylphthalate). After polymerization, the coverslip was removed by quick cooling leaving a flat re-embedded portion of the original section. This portion was viewed by transillumination in a dissecting microscope and trimmed of surplus tissue. Ultrathin sections for electron microscopy were obtained in the usual manner.     

A Controllable Carmine Technic for Plants with Small Chromosomes

Anthers of small chromosome plants (Antirrhinum, Brassica, Capsicum etc.) were fixed 12 hours or longer at 0-3° C. in: ferric acetate in glacial acetic acid (sat. soln.), 1 part; absolute alcohol, 3 parts. They were transferred to: ferric acetate (sat. soln.) in 45% acetic acid, 3 parts; 45% acetic acid, 5 parts; 1% formalin (aq.), 2 parts, and allowed to remain 5-15 minutes at room temperature for mordanting. The amount of iron introduced into the specimens was controllable by the time in the mordanting fluid. After rinsing the specimen in 45% acetic acid and macerating in a drop of Belling's acetocarmine on a slide, a cover slip was applied followed by warming and pressing with blotting paper to flatten the pollen mother cells and expel excess stain. Preparations stored temporarily by sealing the edges of the cover slip with rubber solution were best made permanent by removing the cover slip after 1-2 days, dehydrating and mounting in euparal.     

Azure a-Schiff, Alcian Blue, HIO4-Schiff, Naphthol Yellow S—A Sequential Staining Method for Paraffin Sections

Paraffin sections from tissue fixed 4-12 hr in 10% formalin containing 0.5% cetyl pyridinium chloride, and washed 2 hr, were stained as follows: (1) Hydrolyze in 5 N HCl at room temperature for 8.5-9 min, or use standard Feulgen hydrolysis at 60 C. (2) Stain in azure A-Schiff, 0.5% in bisulfite bleach (1 N HCl, 5; 10% Na₂S₂O₅, 5; and distilled water 90—parts by volume) for 10 min. (3) Place in bisulfite bleach 2 changes, 2 min each; wash in water, 1-2 min. (4) Stain in Alcian blue (0.1% in 0.01 2V HCl, pH 2.0) for 10 min. (5) Place in 0.01 N HCl for 2-3 min; wash in water for 1-2 min. (6) Oxidize in 0.5% HIO₄ for 5 min; wash in water, 1-2 min. (7) Stain in Schiff's leucofuchsiu, 10 min. (8) Treat with bisulfite bleach as in step 3; wash in running water, 10 min. (9) Stain in naphthol yellow S (0.01% in 1% acetic acid) for 1-2 min. (10) Place in 1% acetic acid for 2 min, dehydrate in tertiary butanol, clear and cover. Result: DNA is deep blue; acidic mucins are light blue; neutral polysaccharides, red to magenta; and proteins, yellow. Proper timing of the hydrolysis for the Feulgen reaction is the most critical step. Overhydrolysis results in green nuclei (staining by naphthol yellow S) whereas purplish nuclei are the results of insufficient hydrolysis.