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.     

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.     

An Acetic Acid Dissociation, Air-Drying Technique for Insect Chromosomes, With Aceto-Lactic Orcein Staining

The method differs from mammalian techniques for somatic chromosomes in that it uses very small amounts of material. Drosophila melanogaster and an ant, Dorymyrmex sp., are used as examples. Pretreatment with 0.05% Colcemid in insect Ringer solution is applied to mature Drosophila larvae for 5 hr, by feeding, but Dorymyrmex prepupae require puncture and a 15 hr exposure of the puncture to the solution. Organs are removed under 1% sodium citrate, tansferred to fresh citrate for 10-20 min, than fixed in acetic-methanol, 1:3, for 30 min. Transfer to a drop of 60% acetic acid on a clean warmed slide dissociates the cells, which are spread by adding a small drop of fixative and tilting the slide in all directions. After immersion in acetic ethanol, 1:3, for 4 hr, rinsing in the stain solvent and draining the slides then have 2-3 drops of aceto-lactic orcein placed on each, coverslips added, and warmed (at about 50 C) for about 12 hr or until staining is sufficient. They can then either be treated as semipermanent or made permanent by allowing the coverslips to slide off in acetic-ethanol, dehydrating, and mounting in Euparal, or a synthetic resin.     

Reversible, positive cooperative interaction of 11 beta-chloromethyl-[3H]estradiol-17 beta with the calf uterine estrogen receptor

The binding of 11 beta-chloromethyl-[3H]estradiol-17 beta [3H]CME2) with the calf uterine estrogen receptor was investigated. The equilibrium binding analysis indicated a positive cooperative interaction yielding curvilinear Scatchard plots and Hill coefficients of 1.4-1.5. This positive cooperative interaction of [3H]CME2 was indistinguishable from the typical cooperative interaction of [3H]estradiol with the receptor. The apparent relative association constant and the relative binding affinity of CME2 for the estrogen receptor measured by competitive binding assay were 146 and 184%, respectively. The dissociation kinetics of [3H]CME2 from the receptor was biphasic, composed of a fast dissociating component (15%, t1/2 = 4 min at 0 degrees C; 9%, t1/2 = 4 min at 28 degrees C) and a slow dissociating component (85%, t1/2 greater than 50 h at 0 degrees C; 91%, t1/2 greater than 50 h at 28 degrees C). The dissociation kinetics of [3H]estradiol was also biphasic: the t1/2 of the fast dissociating component was 4 min at 0 and 28 degrees C and approximately 200 min for the slow dissociating component at both temperatures. The fraction of the slow [3H]estradiol dissociating component increased from 56 to 92% upon warming. Ethanol extraction and trichloroacetic acid treatment proved that the binding of [3H]CME2 is fully reversible. The unusual dissociation kinetics and the binding mechanism of CME2 are discussed.     

Staining and Mounting Helminths

The following procedure is recommended: Fix ces-todes and trematodes (while held flat between glass slides) 0.5-2.0 hr. in the following mixture: formalin, 15; acetic acid (gl.), 5; glycerol, 10; 95% ethyl alcohol, 24; distilled H₂O, 46; all proportions by volume. After freeing them from the slides, wash thoroughly in running water and stain immediately thereafter. Stock staining solution: ferric ammonium alum (violet cryst.), 2 g.; distilled H₂O (cold) 100 ml.; after solution, add 2 ml. concentrated H₂SO₄, bring to a boil; add 1 g. coelestin blue B (Nat. Aniline), boil 3-5 min.; cool and add 10 ml. absolute methyl alcohol and 10 ml. glycerol. Dilute 1 vol. with 3 vol. distilled H20 for use. Stain 5-30 min., depending on size of specimens. Wash with 2 changes 0.5 hr. each of distilled H₂O, then 50% isopropyl alcohol 12-16 hr., 50% isopropyl alcohol 2 hr., followed by graded isopropyl alcohol for dehydration. Ether: ethyl alcohol (equal parts), 1 hr., is followed by embedding in celloidin in a sheet just thick enough to cover the specimens. Trim embedded specimens and dehydrate with isopropyl alcohol, 80%, 90% and absolute. Clear in beechwood creosote. Mount in balsam with cover glasses that overlap the edges of the celloidin 1-2 mm. While drying at 37°C, refill edges of mount with fresh balsam as needed. When dry, remove excess balsam and ring the edges with ordinary gloss enamel paint.     

A Method for the Selective Removal of Deoxyribonucleic Acid from Tissue Sections

The deoxyrihonucleic acid (DNA) of chromatin undergoar depurinization on mild acid hydrolysis with a picric acid-formaldehyde mixture (Bouin's fluid). The apurinic acid thus formed is degraded by condensation with aniline and is lost from tissue sections, but ribonucleic acid (RNA) in nucleoli and cytoplasm is well preserved. Technique: Fi in Carnoy's fluid (ethanol:acetic acid 3:1 or ethanol:chloroform:acetic acid 6:3:1) or in aldehydes (10% formalin or 2.5% glutaraldehyde bsered to pH 7.0). Hydrolyse deparaEnii sections 12-24 hr at 27-50 C in Bouin's fluid, wash in distilled water, immerse in 25% (v/v) acetic acid, treat 1 hr at 27-30 C with 10% (v/v) dine in 25% acetic acid, wash in 25% acetic acid and then in water. Stain 10-40 min with 0₃% toluidine blue in 0.05 M potassium biphthalate bder (pH 4.0); rinse in distilled water, pass to 10% (w/v) ammonium molybdate for 1 min, rinse again in water and pass through tert-butanol and xylene to a synthetic resin. Chromatin and chromosomes are pale green; RNA in nucleoli and cytoplasm deep purple.     

Nile Blue Histochemical Method for Phospholipids

The technic recommended is: Fix 6-12 hr. in 10% formalin containing 1% CaCl₂. Cut frozen sections without embedding or after gelatin or carbowax. Stain 90 min. at 60°C. in saturated aqueous Nile blue sulfate, 500 ml. plus 50 ml. of 0.5% H₂SO₄, boiled 2 hr. before use. Rinse in distilled water, and place in acetone heated to 50°C. Remove the acetone from the source of heat and allow the sections to remain 30 min. Differentiate in 5% acetic acid 30 min., rinse in distilled water, and refine the differentiation in 0.5% HCl for 3 min. Wash in several changes of distilled water and mount in glycerol jelly. Results: phospholipids - blue; everything else - unstained. Counterstaining nuclei with safranin is optional, but if done, it preferably precedes the Nile blue and is then differentiated by the acetic acid. The histochemical principles on which the method is based are as follows: (1) The calcium compounds of phospholipids combine with the oxazine form of Nile blue sulfate and survive subsequent treatment; (2) neutral lipids are dissolved out by acetone; (3) proteins and other interfering substances are destained by the acetic acid and hydrochloric acid baths.     

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.     

Direct and Repeatable Bone Marrow Chromosome Preparations from Living Mice

Using a 27 gauge hypodermic needle, bone marrow is aspirated from a lumbar vertebra into 0.1 ml of Hanks' salt solution. The aspirate is kept well mixed in 1% sodium citrate for 15 min, centrifuged, and the cell pellet fixed for 30 min in Clarke's 3:1 ethanol-acetic fixative. After removal of the fixative the cells are suspended in 0.05-0.1 ml of 60% acetic acid, centrifuged and resuspended in 0.03 ml of this fixative. Chromosome preparations are made by spreading the suspension on a slide heated to 60 C.     

Toluidine Blue-Alizarin Red S Staining of Cartilage and Bone in Whole-Mount Skeletons in Vitro

Cartilage and bone of the developing skeleton can be reliably differentiated in whole-mount preparations with toluidine blue-alizarin red S staining after FAA fixation. The recommended staining procedure is based chiefly on the use of newborn white and Swiss-Webster mice, 4-9 days postnatal, but was tested also on mice and rats 3-8 wk of age. Procedure: Sacrifice, skin, eviscerate, remove body fat, and place specimens in FAA (formalin, 1; acetic acid, 1; 70% alcohol, 8) for approximately 40 min. Stain in 0.06% toluidine blue made in 70% ethyl alcohol for 48 hr at room temperature. Use 20 volumes of stain solution to the estimated volume of the specimen. Destain soft tissues in 35% ethyl alcohol, 20 hr; 50%, 28 hr; and 70%, 8 hr. Counterstain in a freshly prepared 1% aqueous solution of KOH to which is added 2-3 drops of 0.1% alizarin red S per 100 ml of solution. Each day for 3 days, transfer the specimen to a fresh 1% KOH-alizarin mixture, or until the bones have reached the desired intensity of red and soft tissues have cleared. Rinse in water, and place in a 1:1 mixture of glycerol and ethyl alcohol for 1-2 hr, then transfer the specimen to fresh glycerol-alcohol for final clearing and storage. Older mice and rats require procedural modifications: (1) fixation for 2 hr, (2) 0.12% toluidine blue, (3) maceration for 4 days in 3% KOH-alizarin, and (4) preliminary clearing for 24 hr in a mixture of glycerol, 2; 70% ethyl alcohol, 2; and benzyl alcohol, 1 (v/v) before placing in a 1:1 alcohol-glycerol mixture.     

Fine cytogenetical analysis of the band 10a1-2 and the adjoining regions in the Drosophila melanogaster X chromosome

The region 9E1-2 - 10B1-2 of the Drosophila melanogaster X chromosome was analysed under the light (LM) and the electron (EM) microscope using different fixatives and an EM map of the region was constructed. EM analysis revealed 21 bands in the region 9E1-2 - 10B1-2 instead of 36 bands in Bridges' map. This discrepancy mainly results from the fact that 14 bands indicated as "doublets" by Bridges appear as a single bands. No doublets were found in the whole 9B1-2 - 10C1-2 region after fixation of salivary glands in 3% glutaraldehyde, 3% formaldehyde and 3 : 1 ethanol-acetic acid mixture. 45% acetic acid is the only fixative which results in strongly vacuolated appearance of the bands. - The break points of 30 chromosome rearrangements in the region 9E1-2 - 10B1-2 were located under EM or LM within the limits of the EM map of this region.     

Effect of Fixatives on Silver Impregnation of Plant Cells

The kind of fixative and duration of fixation modify the affinity of plant cell structures, as shown by a 10-15 hr impregnation at 70 C in 2% aqueous AgNO₂, and a 1-2 hr reduction at room temperature by a 1:1 mixture of 10% formalin and 1% hydroquinone. Cytoplasmic staining was enhanced by fixing in salts of heavy metals, in buffered 6.5% glutaraldehyde, and in 0.5% picric acid. Nuclear staining was prominent after mixtures of glutaraldehyde and hydroquinone, after formalin and pyrogallol, and after acetone, propylene glycol or ether. Nucleolar staining was favored by fixing in 10% formalin, in 5% formalin containing 0.5% hydroquinone, in 50% ethanol containing 0.5% pyrogallol, or in ethylene glycol. Chromosome staining was favored by fixation in 50% acetic or propionic acid, in 2% trichloroacetic acid, and in methanol or ethanol. The best morphological preservations were seen after 50% acetic acid, 6.5% glutaraldehyde, or the 5% formalin-0.5% hydroquinone mixture.     

A Sensitive Myelin-Sheath Stain Particularly Useful for Small Mammals and Neonatal Cats

Staining of myelinated fibers including the delicate myelin sheaths of infantile animals is as follows: perfuse the anesthetized animal with a pH 7.4 posphate-buffered fixative, either 10% formalin, 6% gluteraldehyde or a mixture containing 3% gluteraldehyde and 2% acrolein. Dissect out the brain or spinal cord and continue fixation for at least 24 hr. Cut larger brains to 1 cm in at least one dimension. Wash in running tap water 2-3 hr and soak in 2.5% potassium dichromate in 1% acetic acid (the primary mordant) for 3-5 days in darkness. Wash at least 12 hr in running tap water. Dehydrate and embed in celloidin and store in 80% ethanol. Section at 25-60 μ into 80% ethanol. Wash 1-2 min in distilled water and then immerse in 1-2% ferric alum at 50 C for at least 1 hr (the secondary mordant). Wash in tap water and stain at least 1 hr at 50-60 C in 0.5% unripened hematoxylin in 1% acetic acid. Wash well in tap water and differentiate in a mixture containing 0.5% ferrityanide, 0.5% borax and 0.5% Na₂CO₃; 2 changes. Wash well in distilled water, then in tap water, and dehydrate, clear and mount. Myelin stains black, cell bodies stain tan, and the background is pale yellow. With minor modifications in timing, the method is applicable to frozen and to paraffin sections; the primary mordant being omitted in the freezing technique.     

Slide Processing for the Examination of Male Mammalian Meiotic Chromosomes

A rapid method was devised specifically for the cytological identification of translocations in the male mouse at late prophase to metaphase of meiotic division I, but the method should be useful for less specific objectives requiring examination of mammalian testicular material. For the adult mouse, masses of tubules from a single testis, freed of the testicular tunic, are placed in 3 ml of 0.7% sodium citrate for 15-20 min, and subsequently fixed in 50% acetic acid by the addition of 3 ml of glacial acetic acid to the hypotonic citrate. To facilitate handling of individual tubules by preserving their visible structure, the addition of fixative is at a rate which is grossly adjusted so that 2 ml will have been added at the end of 30 sec and the remaining 1 ml by the end of a minute. A single fixed tubule 1-2 cm long is placed lengthwise on a slide and covered with a drop of lactic-acetic orcein made as follows: Add 2 gm of synthetic orcein (G. T. Gurr) to a mixture of 50.0 ml of glacial acetic acid, 42.5 ml of 85% lactic acid, and 7.5 ml of distilled water. After staining for 10 min, a 22 × 50 mm cover slip is placed over the tubule, and it is allowed to stain for an additional 10 min. The majority of germinal cells will not be in late prophase or metaphase of the first meiotic division, therefore many preparations will be useless; however, slides with division figures are radidly selected as follows: Before squashing, examine under a microscope at a magnification of 150, and upon recognition of a single meiotic division, remove the slide and squash the preparation for subsequent detailed examination. As a consequence of the spermatogenic wave that progresses along the length of a tubule, a given slide will usually have many division figures or none at all, hence the limitation of 1 tubule per slide facilitates efficient discarding. Preliminary work with the Chinese hamster suggests that good preparations might be obtained from testes of various mammals when the volume of hypotonic solution is adjusted so as to compensate for varying testicular sizes by maintaining a 15:1 ratio of fluid to estimated volume of tissue.     

An improved method for preparing whole specimens from bovine preimplantation embryos: A technique note

A method was devised to prevent loss of whole embryos during fixation. Specimens were prepared in a chamber saturated with fixative vapors consisting of 3 : 1 (v/v) 96%. ethanol/glacial acetic acid. Good quality specimens were obtained after fixation for at least 24 but not more than 72 h. After staining, specimens could be preserved for 3 to 4 d by storage in the fixation chamber, in 45% aqueous acetic acid vapor. Using the method suggested in this paper prevents loss of early embryos during fixation and allows storage of specimens for longer than usual time while maintaining the quality of the specimen.     

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.     

Anaerobic conversion of lactic acid to acetic acid and 1, 2-propanediol by Lactobacillus buchneri

The degradation of lactic acid under anoxic conditions was studied in several strains of Lactobacillus buchneri and in close relatives such as Lactobacillus parabuchneri, Lactobacillus kefir, and Lactobacillus hilgardii. Of these lactobacilli, L. buchneri and L. parabuchneri were able to degrade lactic acid under anoxic conditions, without requiring an external electron acceptor. Each mole of lactic acid was converted into approximately 0.5 mol of acetic acid, 0.5 mol of 1,2-propanediol, and traces of ethanol. Based on stoichiometry studies and the high levels of NAD-linked 1, 2-propanediol-dependent oxidoreductase (530 to 790 nmol min(-1) mg of protein(-1)), a novel pathway for anaerobic lactic acid degradation is proposed. The anaerobic degradation of lactic acid by L. buchneri does not support cell growth and is pH dependent. Acidic conditions are needed to induce the lactic-acid-degrading capacity of the cells and to maintain the lactic-acid-degrading activity. At a pH above 5.8 hardly any lactic acid degradation was observed. The exact function of anaerobic lactic acid degradation by L. buchneri is not certain, but some results indicate that it plays a role in maintaining cell viability.     

A Versatile Stain for Pollen Fungi, Yeast and Bacteria

A versatile stain has been developed for demonstrating pollen, fungal hyphae and spores, bacteria and yeasts. The mixture is made by compounding in the following order: ethanol, 20 ml; 1% malachite green in 95% ethanol, 2 ml; distilled water, 50 ml; glycerol, 40 ml; acid fuchsin 1% in distilled water, 10 ml; phenol, 5 g and lactic acid, 1-6 ml. A solution has also been formulated to destain overstained pollen mounts. Ideally, aborted pollen grains are stained green and nonaborted ones crimson red. Fungal hyphae and spores take a bluish purple color and host tissues green. Fungi, bacteria and yeasts are stained purple to red. The concentration of lactic acid in the stain mixture plays an important role in the differential staining of pollen. For staining fungi, bacteria and yeasts, the stain has to be acidic, but its concentration is not critical except for bacteria. In the case of pollen, staining can be done in a drop of stain on a slide or in a few drops of stain in a vial. Pollen stained in the vial can be used immediately or stored for later use. Staining is hastened by lightly flaming the slides or by storing at 55±2 C for 24 hr. Bacteria and yeasts are fixed on the slide in the usual manner and then stained. The stock solution is durable, the staining mixture is very stable and the color of the mounted specimens does not fade on prolonged storage. Slides are semipermanent and it is not necessary to ring the coverslip provided 1-2 drops of stain are added if air bubbles appear below the coverslip. The use of differentially stained pollen mounts in image analyzers for automatic counting and recording of aborted and nonaborted pollen is also discussed.     

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.     

Aceto-Iron-Haematoxylin-Chloral Hydrate for Chromosome Staining

Aceto-iron-haematoxylin can be used combined with the clearing agent chloral hydrate for the squash method. The stain is prepared by dissolving 2 gm of chloral hydrate in 5 ml of a stock solution of 4% haematoxylin and 1% iron alum in 45% acetic acid, which has been allowed to ripen for 24 hr to 1 wk. Heat must not be used to hasten solution. The material (fixed in 1:3 acetic-alcohol) is put on a slide, the fixative removed and a drop of stain added; if necessary the material is crushed before the cover slip is placed in position. The preparations are now carefully heated until a slight colour change occurs. Squashing needs more pressure than in other techniques. This stain gives best results in zoological and botanical material not requiring hydrolysis, e.g., leucocytes, ascites cells, and cells undergoing spermatogenesis and microsporogenesis. Well-spread and selectively stained mitotic and meiotic figures can be obtained.