A Modified Bradley's Squash Technique for Studying Embryo Sacs in Gramineae

This procedure is especially suited for studying the embroyology of sexual and apomictic grasses. Material is fixed in a 2:2:1 alcohol-chloroform-propionic acid mixture for a minimum period of 2 days, soaked in 4% iron alum at 75 C for 7 min, and 2 min each in 2 changes of distilled water, also at 75 C. After 2-3 min in cold water, it is macerated in 50% HCI for 10 min at about 22-25 C, washed and mordanted for 12-16 hr in 50% alcohol saturated with ferric acetate. Ovules are then dissected out and squashed in 1% carmine in 45% propionic acid. Squashing should be firm enough to separate and flatten the embryo sacs but not to burst them. The slides are set aside for 12-24 hr for intensification of the stain.     

Cold and Chemical Pretreatments to aid Chromosome Counts in a Grass Leaf Squash Technique Incorporating Hot Pectinase Maceration

Leaf buds, comprising the basal 3-5 mm of the youngest leaves attached to short stems, were dissected out of fast-growing young tillers of certain grasses, including Festuca, Lolium and Phalaris spp. and various hybrids. They were kept overnight in distilled water at 0-2 C, treated in a mixture of equal parts by volume of saturated aqueous solutions of 5,7-dibromo-8-hydroxyquinoline containing a surfactant (Tween 80), and 1-bromo-naphthalene for 3-4 hr at 0-2 C, and fixed in Newcomer's fluid. The rinsed samples were hydrolysed in 1 N HCl for 8 min at 60 C and Feulgen stained for 1 hr. After rinsing, the buds were macerated in a filtered 3% solution of Pectinol 100-D (Rohm and Haas) in 0.1 M acetate buffer at pH 4.5 for 10 min at 60 C. Squashes were made in 45% acetic acid. The combined cold and chemical pretreatments resulted in strongly contracted, easily counted metaphase chromosomes, while intact cells with full chromosome complements were more readily retained during squashing after enzyme maceration at 60 C than at room temperature.     

Adaptation of the Millon, Sudan Black B, and Periodic Acid-Schiff Technics for Block Staining of Insect Tissues

Spermatophores and reproductive systems of the beetle, Lytta nuttalli Say, fixed in Bouin's aqueous picroformol or buffered 10% neutral formol were stained in toto by the Millon, Sudan black B and periodic acid-Schiff reactions as follows. Millon: after excess fixative is removed in 70% ethanol, specimens are brought to water, stained in Millon's reagent at 60 C for 1 hr, rinsed in 2% aqueous nitric acid at 40-50 C, dehydrated rapidly, cleared, embedded and sectioned as usual. Sudan black B: specimens are taken to absolute ethanol, stained in a saturated solution of Sudan black B in absolute ethanol at room temperature for 24-48 hr, rinsed and cleared in xylene, embedded and sectioned. PAS: specimens are brought to water, oxidized in 0.5 aqueous HIO₄ at 37 C for 30 min, washed in 2 changes of water, stained in Schiif reagent at room temperature for 1 hr, rinsed in 3 changes of 0.5% aqueous potassium metabisulfite, washed in running water for 10-15 min, dehydrated, cleared, embedded and sectioned. All 3 methods produced their characteristic staining in specimens up to 3 mm thick     

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.     

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.     

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 Combined Hcl-Acetic-Alcohol Fixation and Hydrolysis Followed by Cresyl Violet Staining for Mosquito Chromosome Spreads

Mosquito tissues of cytogenetical importance were dissected out on a slide in 0.65% NaCl, under a dissecting microscope, and treated about 30 sec in a drop of 1:3 Carnoy's fixative diluted 1:19 with distilled water. Fixing and hydrolysis was done by a single step in a mixture consisting of: glacial acetic acid, 1; ethanol 96%, 3; HCl conc., 2; and distilled water, 2 (v/v) for 2-6 min at 20-25 C. The specimen was then rinsed with the acetic-alcohol fixative and covered in a drop of 1% cresyl violet in 50% acetic acid under a coverslip coated with Mayer's albumen. Washing was performed immediately by adding water dropwise to one side of the coverslip and drawing the fluid from the other side with absorbent paper. The preparation could be used either as a temporary slide or made into a durable mount. The DNA-containing bands of the giant polytenic chromosomes stained dark violet; interband regions, weakly stained or colourless against a clear background. Mitotic and meiotic figures in gonadal cells stained selectively dark violet or violet with a practically unstained cytoplasm.     

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.     

A Colchicine, Hypotonic Citrate, Air Drying Sequence for Foetal Mammalian Chromosomes

Mice 14 days pregnant were given 0.3 ml of 0.025% Colcemid (Demecolcine, Ciba) and killed after 1 hr. The livers of the foetuses were removed and broken up in 0.1% Colcemid in phosphate-buffered 0.85% NaCl and left 1.5 hr. The cell suspension was then centrifuged, resuspended in 1% sodium citrate for 20 min, centrifuged and the cells fixed in acetic-alcohol (1:3) for 30 min at 4°C. The cells were then resuspended (twice) in 45% acetic acid and dropped onto warm slides. After drying, the cells were stained for 30 min with lactic-acetic-orcein and examined under oil-immersion without a cover slip. Good numbers of well-spread mitotic figures were obtained.     

Double Staining of Plant Materials in Bulk

Ovaries and ovules of Oryza sativa and Zea mays were collected between 9-30 and 10-30 AM, fixed in formalin-acetic-alcohol, stained in Delafield's hematoxylin for 2-4 hr, dehydrated through graded ethanol, counterstained for 3-4 hr either in light green, orange G or fast green (0.05-0.1%) at the 1:1 alcohol-xylene stage and embedded. A few ovaries were hydrolysed in 1 N HCI for 25 min at 60 C, stained in leuco basic fuchsin for 60-90 min, rinsed 3 times with a mixture of: 10% Na₂S₂O₅, 1; N HC1, 1; and distilled water, 18; washed repeatedly in distilled water, dehydrated through graded ethanol, counter-stained for 3-4 hr either with light or fast green (0.05-0.1%) at the 1:1 alcohol-xylene stage and embedded. Microtome sections were cut, ribbons mounted, dried, paraffin removed with xylene, and mounted in balsam. Uniformly stained preparations resulted and the dilute stains gave vivid color contrasts. Large numbers of ovules and ovaries can be processed in a short time, and reliable percentages of viable embryo sacs in normal, sterile and semisterile plants obtained.     

Staining Residual Lipids in Ultrathin Sections of Tissues Embedded in Polyester Resin

Rat suprarenal glands fixed in Palade's 1% OsO₄, buffered at pH 7.7 with veronal-acetate, to which 0.1% MgCl₂ was added, were embedded in Vestopal-W and sectioned at 0.2-1 µ. The sections were attached to slides by floating on water, without adhesive, and drying at 60-80° C, placed in acetone for 1 min and then treated with the following staining procedure: Place the preparation in a filtered solution of oil red O, 1 gm; 70% alcohol, 50 ml; and acetone, C.P., 50 ml; for 0.5-1 hr. Rinse in absolute ethyl alcohol; drain; counterstain with 0.5% aqueous thionin for 5 min; rinse in distilled water; drain; stain in 0.2% azure B in phosphate buffer at pH 9, for 5 min. Dry and apply a drop of immersion oil directly on the section. The preparations are temporary. Ciaccio-positive lipids, rendered insoluble by OsO, fixation, stained red to ochre.     

Serial Sectioning of Insects with Hard Exoskeleton by Dissolution of the Exocuticle

The method reported here was designed to produce paraffin serial sections as thin as 5 Mm of insects or other arthropods with a hard cuticle. Heads and abdomens of Apis mellifera, Eristalomyia tenax and Tenebrio molitor were fixed with Schaffer's liquid, dehydrated with 80% ethanol, 90% ethanol, two changes of 100% isopropanol (2 hr each) and 12 hr in a 1:1 mixture of paraffin (58 C melting point) at 60 C. They were molded in paraffin after 12 hr of infiltration under a partial vacuum at 60 C. Large body openings of objects were sealed with paraffin to prevent infiltration of solvents.

Thereafter, the outer paraffin was removed manually and with xylene (15 min); the cuticle was rehydrated with 100% isopropanol and 100% ethanol (15 min each). The objects were then treated with Sputofluol (Merck; a mixture of NaOH and NaCIO) until they became white or their colorless endocuticle was stainable with aniline blue WS (C.I. 42755) after rinsing in a 50% acetic acid solution (v/v). They were then dehydrated with 100% ethanol and 100% isopropanol (15 min each) and subsequently re-embedded in paraffin. They were molded, sectioned, stained and mounted as usual.     

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).     

Display of mitochondria in root tip cells

A method for displaying mitochondria and proplastids in root tip sections of Tradescantia paludosa and cereals was modified from Altmann and Volkonsky. Root tips were fixed in 3% glutaraldehyde in phosphate buffer, pH 7.1, or acetate buffer, pH 4.8, for 3 hr, rinsed and postchromed overnight in 3% potassium dichromate, all at room temperature (20 C), dehydrated through a tertiary butanol series and embedded in ester wax. Four-micrometer sections were stained in hot acid fuchsin in aniline water, rinsed, treated with 1% sodium phosphomolybdate for 30 sec, rinsed and stained progressively with azure B for 3-10 min before being made permanent. Mitochondria and proplastids were stained brilliant crimson against a light blue cytoplasm with deep blue chromosomes. Previously reported difficulties with Altmann staining techniques are attributed to the erratic action of the classical fixatives used.     

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.     

Camwood (Baphia nitida) Extract as a Histological Stain for Interglobular Dentine, Striated Muscle, and Counterstaining

The shavings of the dried heartwood of the tree Baphia nitida are ground to a fine powder, and 6 gm of the powder are extracted in 100 ml absolute ethanol at 27-30 for 6-24 hr. The extract is filtered with Whatman No. 1 paper and stored in a screw-capped bottle. For staining the interglobular dentine of nondecalcified sections of formlin-fixed teeth, sawed cross sections 20-30 μ thick were dehydrated in ethanol and stained in the undiluted extract for 6-12 hr at room temperature. The interglobular dentine was stained a bright golden brown on a pale brown background. For staining striated muscle, the extract was diluted 1:1 with distilled water and filtered. After mordanting formalin-fixed paraffin sections with 0.25% KMnO₄ for 5 min, and bleaching with 5% oxalic acid for 10 min, they were washed in water and stained for 2-24 hr at room temperature. The striations were stained light to deep golden brown. For use as a counterstain, a 1:6 dilution of the original extract was required. When applied after haematoxylin for 15-30 min, it stained tissue components in varying shades of golden brown with distribution comparable to that produced by 1% eosin.     

The influence of formaldehyde fixation time on the rate of nitrous acid deamination of erythrocytes and spinal cord proteins

Summary Alcohol fixed blood films and fresh blocks of spinal cord were immersed in phosphate buffered neutral 10% formol for graded intervals, the films for 10, 30 min, 1, 2, 4, 8, 24 hr; the blocks for 2, 4, 6, 24 hr at 3 and 24° C; 1, 3, 7, 14, 21, 28, 42, 56 da, 3 and 14 mo at 24–26°. Graded deaminations in 2 N NaNO₂/HAc at 3° C were applied: 1, 2, 5, 10, 20, 30 min; 1, 2, 4, 6, 8, 12, 18, 24, 36 hr. Blood films were stained at pH 6 and 6.5, tissue at pH 4.5 and 5.0, both in azure A eosin B. The point at which erythrocytes reached a slightly bluish green was taken as the end point, since no further color change occurred on further exposure and erythrocytes were the last of usually deamination susceptible tissue elements to lose their oxyphilia on deamination. Deamination of alcohol fixed blood films is completed in about 2 min, of sublimate fixed spinal cord in about 1 hr. Progressive formaldehyde exposure increased deamination time of blood films to 10–20 min in 1 hr, to 6–8 hr in 4 hr and to 12 hr in 24 hr. The tissue deamination showed similar progressive increase of deamination time, slower with 3° C fixation than with 24–26°, reaching 18–36 hr by about 3 days formol, and remaining about the same thereafter.Supported by National Cancer Institute Grant No. C-04816, National Institutes of Health.     

Silver Staining of Ultrathin Sections of Tissues Embedded in Polyester Resin

Specimens 1 mm³ from rat liver and kidney were fixed for 50 min in cold (0-2° C) 1% OsO₄ in veronal-acetate buffer, pH 7.7, and containing 0.1% MgCl₂; then dehydrated and embedded in Vestopal-W. Sections were cut in two ranges, 0.1-2 µ and 60-90 mµ thick, and attached to slides by floating on water and drying at 60° C. The thicker ones, for light microscopy, were soaked in acetone 1.5-3 hr; the thinner, for electron microscopy, 20-30 min. Both kinds were stained by Wilder's (1935) method for reticulum. Those for light microscopy were finished by dehydrating, clearing and covering in the customary manner; those for electron microscopy, by coating with 1% parlodion, drying, cutting the film about 2 mm² around the section, and freeing the section by soaking in water. The section was then mounted on a grid. The structures stained are: nuclei, basement membrane of capillaries, reticulum fibers of the liver and kidney, and in addition, the basement membrane of the kidney tubules. The mitochondria, vesicles, endoplasmic reticulum and cell membranes were not defined.     

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 Osteoid Seams in Thin Slabs of Undecalcified Trabecular Bone

For qualitative and quantitative study of osteoid seams in trabecular bone, 5-7 mm thick slabs were cut from the bodies of fresh, frozen, undecalcified human vertebrae. After washing out the bone marrow and soft tissue in a jet stream of water, the slabs were stained in 0.5% aqueous basic fuchsin for 30-40 hr at 18-20 C. The specimens were then trimmed by sawing off both overstained surfaces, to make a 2 mm slab which was submerged in 50% ethanol (2 or 3 changes of 10-30 min each), until the nonosteoid trabeculae became pale pink. The slab was allowed to dry in air. Osteoid seams are stained dark red and are well differentiated under a dissecting microscope with reflected illumination, either dry or immersed in water. This method permits the various types of trabculae to be separately studied in the same specimen