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).     

Removal of Ribonucleic Acid from Bacterial Cells by Dilute Salt Solutions

Bacterial cells were impressed upon a clean glass slide, fixed in ethyl alcohol and immersed at 37°C in either of the following two salt solutions: (A) NaCl, 7.8 gm; KCl, 0.7 gm; distilled water, 1000 ml; adjusted to pH 7.0; or (B) 0.1M NaH₂PO₄, 400 ml; 0.1M Na₂HPO₄, 600 ml; KCl, 0.7 gm. After 1-5 hr soaking to remove ribonucleic acid, the slide was stained by Giemsa's method as usual. The staining revealed slender chromatinic bodies with reasonable clarity extending the whole diameter of the moderately swollen cell. The results of this method seemed to be much like those obtained after ribonuclease digestion.     

Application of Methods for Mammalian Chromosome Spreads to the Cells of Cestodes

Tapeworm cells obtained by physical maceration between ground-glass surfaces are incubated for 3 hr in Hanks' balanced salt solution (BSS) supplemented with colchicine to a concentration of 10-⁴ M. After washing in BSS, the cells are incubated for 10 min in 1/4 strength BSS then centrifuged 10 min. Fixation of the intact button of cells (or alternatively, by squirting the cells directly into the fixative) in Carnoy's alcohol-chloroform-acetic acid (6;3:1) for 30 min follows, and cells, dispersed and washed in the fixative, are flattened by dropping the suspension on clean, water-wet slides which are then air-dried and stained with Giemsa diluted 1 ml;47 ml with distilled water to which 2 ml of buffer—M/15 KH₂PO₄, 32 ml, mixed with M/15 Na₂HPO₄, 68 ml—is added. After staining 15 min and washing in distilled water, slides are air-dried and mounted with resin. Well separated and well stained chromosomes have resulted.     

Plastic Sectioning for Light Microscopy of Pyrenomycetes

In preparation for light microscopy, ascocarps of Sordaria fimicola Ces. & DeNot. were embedded in Spurr's medium and sectioned at 1-1.5 μm on an ultramicrotome. Sections were floated on Giemsa staining solution at 60 C for 10-30 min, washed in distilled water, affixed to slides by drying, and mounted in immersion oil. Best preservation of the delicate sterile tissues of the centrum was obtained by fixation in 1% KMnO₄ for 2.5-3 hr, followed by the Giemsa stain. This method is suggested for future studies on the morphology of perithecial ascomycetes.     

Staining of fresh, Undecalcified, thin Bone Sections

Fresh, undecalcified bone sections can be reproducibly and reliably stained by any of the following procedures: (A) Basic fuchsin, 1% in 30% alcohol, 48 hr, 22°C. (B) AgNO₃, 0.033 M, 48 hr, 22°C; washing 48 hr in a large volume of distilled water; exposure to light to develop the color. (C) Metallic sulfides (Co⁺⁺, Pb⁺⁺, Hg⁺⁺, Cu⁺⁺): the nitrate of the metal, 0.033 M, 48 hr, 22°C; then Na₂S, 0.033 M, 48 hr, 22° C. (D) Alizarin Red S, 0.1% solution in distilled water, 48 hr, 22°C; differentiated 48 hr at 22°C in weakly alkaline water, pH about 8. (E) KMnO₄: boiling 8-10 min in a 0.1 N, solution. With the exception of D the surface stain must be ground off the section for microscopic examination of its interior. Stain concentration, time and temperature can be altered to suit specific needs.     

Plastic sectioning for light microscopy of Pyrenomycetes.

In preparation for light microscopy, ascocarps of Sordaria fimicola Ces. & DeNot. were embedded in Spurr's medium and sectioned at 1-1.5 mum on an ultramicrotome. Sections were floated on Giemsa staining solution at 60 C for 10-30 min, washed in distilled water, affixed to slides by drying, and mounted in immersion oil. Best preservation of the delicate sterile tissues of the centrum was obtained by fixation in 1% KMnO4 for 2.5-3 hr, followed by the Giemsa stain. This method is suggested for future studies on the morphology of perithecial ascomycetes.     

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.     

Streptococcal Desoxyribonuclease for the Removal of Feulgen-Stainable Material

Removal of Feulgen-stainable material from the cell nucleus was accomplished by treatment of sections with streptococcal desoxyribonuclease. The procedure recommended is (1) Deparaffinize with xylene, followed by descending grades of alcohol. (2) Wash in tap water. (3) Treat slides for 1 hour at 37°C. with streptococcal desoxyribonuclease (1000 units/ml.) in 0.025M veronal buffer of pH 7.5 containing 0.003M MgSO₄. Treat control slides for an equal length of time at the same temperature. Renew the enzyme approximately every 15 minutes. (4) Wash slides briefly in tap or distilled water. (5) Dehydrate, then coat the sections by dipping in a 1% solution of celloidin in alcohol-ether. (6) Subject the preparations to the Feulgen reaction. Control slides showed characteristic nuclear staining; enzyme treated slides did not stain.     

A Rhodocyan Technic for Staining the Anterior Pituitary

A reproducible, one-step, differential staining technic which uses routine formalin-fixed tissue and gives brilliantly contrasting results is produced by incubating sections for 1 hr in a 60° C oven in the following dye mixture: 1% eosin B (CI#771), 8 ml; 1% anilin blue (CI#707), 2 ml; and buffer solution (0.1M citric acid, 1.1 ml; 0.2M Na₂HPO₄, 0.9 ml; distilled water, 28.0 ml) at pH 4.5. No differentiation is necessary. The method can be modified for duodenal enterochromaffin cells and alpha cells of pancreatic islets by adjusting the buffer to pH 3.6 and staining for only 3 min at 60° C.     

Karyological Leaf Squashes in Grasses, Aided by Pectinase Digestion at 45 C

Longitudinal sections of rapidly growing leaf shoots were soaked for 2-4 hr in 0.002 M 8-hydroxyquinoline at 25 C, blotted, and fixed in 3:4:1 ethanol, chloroform, acetic acid. A 30 min maceration at 45 C in a pectinase solution (Pectinal 59-L; Rohm and Haas) softened the material for staining and squashing. Excess pectinase was removed and 1% aceto-carmine stain was applied. After locating and gently tapping the cover clip to disperse the cells, heavy pressure was applied with a No. 9 rubber stopper and the heel of the hand. By the use of this procedure, karyotypes could be constructed in several genera of forage grasses. The karyotype of Paspalum notatum Flugge is illustrated.     

The Peracetic-Orcein-Halmi Stain: A Stain for Connective Tissues

A selective stain useful for the study of connective tissues is described. The stain demonstrates elastic and oxytalan fibers as well as fibrils in mucous connective tissues previously undescribed. Reticular fibers are not stained. The stain may be used on sections that have been fresh frozen or fixed in formalin or ethanol. Sections are deparaffinized, washed in absolute ethanol, oxidized in peracetic acid 30 min, washed in running water, stained in Taenzer-Unna orcein 15 min, 37°C, differentiated in 70% ethanol, washed in running water, stained in Lillie-Mayer alum hematoxylin 4 min, blued in running water, and counterstained 20 sec in a modified Halmi mixture of 100 ml distilled water, 0.2 gm light green SF, 1.0 gm orange G, 0.5 gm phosphotungstic acid and 1.0 ml glacial acetic acid. Sections are rinsed briefly in 0.2% acetic acid in 95% ethanol, dehydrated and mounted.     

Differential Staining Of Tissue In The Block With Picric Acid And The Feulgen Reaction

The authors have found a modification of the Feulgen reaction to be a satisfactory stain for tissue in the block.

Pieces of fresh mammalian tissue not thicker than 5 mm. are fixed for approximately 48 hours at 25° C. in a mixture of equal parts of 5% aqueous sulfosalicylic acid and saturated aqueous picric acid. They are washed for 30 minutes in three ten-minute changes of distilled water and placed in Feulgen's staining solution diluted to one-half strength with distilled water. The staining solution is allowed to act for 24 hours (2 to 3 mm. thick blocks) up to 48 hours for 5 mm. thickness. After staining, the specimens are transferred to a mixture of sodium bisulfite, 0.5 g. and N hydrochloric acid, 5 ml. in' 100 ml. of distilled water. Two changes of IS to 30 min. each in the acid sulfite are given and these are followed by dehydration through 50%, 70% and 95% alcohol. One to two hours are allowed for each change except the last 95%, in which the stained tissue is allowed to remain overnight. The dehydration is completed in two changes of absolute alcohol with subsequent clearing in xylene and embedding in paraffin. Sections may be cut 10 μ or other thickness desired, mounted on slides, paraffin removed, and covered in the usual manner. Nuclei stain reddish violet against a lemon yellow background when the stain is typical. Orange G, 200 mg. per 100 ml. may be added to the fixing fluid if a more polychromatic effect is desired.     

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.     

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.     

Reverse differential staining of sister chromatids after substitution with BUdR and incubation in sodium phosphate solution

Chinese hamster strain cells were cultured in the presence of BUdR and air-dried on slides. The chromosome preparations were incubated in 1 M NaH₂PO₄ at 88 °C for 4–6 min and stained with Giemsa. The reverse type of sister chromatid differential staining occurred, in which unifilarly BUdR-substituted chromatids stained faintly and bifilarly substituted chromatids stained darkly. Feulgen reaction performed on the same chromosomes after removing Giemsa stain showed the same type of differential staining.     

Toluidine Blue Staining of Coccidia in Cultured Animal Cells

Cultured animal cells infected with various species of Eimeria (coccidia) from chickens are washed in Hanks balanced salt solution (HBSS) and fixed in 5% formalin in HBSS. The fixed cultures are washed briefly in distilled water to remove HBSS salts and then dehydrated in a series of mixtures of 40 to 50% ethanol with increasing concentrations of tertiary butanol (TB) and decreasing concentrations of distilled water. Cultures are placed for 1 min in a mixture of 2 parts ethanol :TB (25:75) and 1 part 0.05% toluidine blue O in McIlvaine buffer (pH 6.0), followed by 1 min in 0.05% toluidine blue O in McIlvaine buffer (pH 6.0). The stained cultures are dipped for 1-2 sec in TB, allowed to dry and mounted permanently on slides. Cover-slip cultures fixed and stained by this procedure 8 years ago have not faded or discolored. The alcohol mixtures, formalin in HBSS, stain and buffer can be prepared in large volumes and stored indefinitely. The staining procedure has proven to be rapid and dependable with a variety of cell types in monolayer cultures in research and teaching applications.     

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.     

Use of Gallocyanin as a Myelin Stain for Brain and Spinal Cord

Tissue fixed in 10% formalin, formol saline, CaCO₃ or phosphate buffer neutralized formalin, Baker's formol calcium, Cajal's formol ammonium bromide, formalin-95% ethanol 1:9, formalin-methanol 1:9, Lillie's methanol-chloroform or Salthouse's formol cetyltrimethylammonium bromide was dehydrated and embedded in paraffin. Sections were attached to slides with either albumen or gelatine adhesive and processed throughout at room temperature of 22-25 C. Mordanting 30-60 min in 1% iron alum was followed by a 10 min wash in 4 changes of distilled water. Myelin was stained in a gallocyanin self-differentiating solution for 1-2.5 hr; thick sections requiring the longer time. The staining solution (pH approximately 7.4) consisted of Na₂CO₃, 90 mg; distilled water, 100 ml; gallocyanin, 250 mg; and ethanol, 5 ml. The ethanol was added to this mixture last, and after the other ingredients had been boiled and then cooled to room temperature. After a staining and thorough washing, Nissl granules were stained for 5-10 min in a solution consisting of: 0.1 M acetic acid, 60 ml; 0.1 M sodium acetate, 40 ml; methyl green, 500 mg. Washing, dehydration, clearing and mounting completed the process. Myelin sheaths were stained dark violet; neuronal nuclei, light green with dark granules of chromatin; nucleoli of motor cells and erythrocytes, dark violet; cytoplasm, green with dark green Nissl granules. The simple and reliable method can be adapted easily for use with automatic tissue processors.