Staining Pericellular Boutons of Glutaraldehyde-Fixed Nervous Tissue; A Chromation-Silvering Sequence for Frozen Sections

After fixing in phosphate-buffered 5% glutaraldehyde, pH 6.8, by perfusion, brains were sliced to 3-5 mm pieces which were placed in the fixative for 5-7 days. The pieces were washed through several changes of 2.26% NaH₂PO₄ for 12 hr, 30 μ frozen sections cut, and mordanted 2 days in an equal-parts mixture of 3.5% CrO₃ and 5% Na-tartrate, which had been aged at 20-25 C for 20 days prior to use. After washing in distilled water, the sections were put into a solution containing AgNO₃, 20 gm; and KNO₃, 15 gm, in distilled water, 80 ml; at 30 C for 1.5-2 hr, then reduced at 40-45 C in three pyrogallol solutions as follows: 1-2 sec in 1% pyrogallol in 55% alcohol; 3-4 sec in a 0.67% solution in 33% alcohol, and 5-7 sec in a 0.5% solution in 25% alcohol. Gold toning is optional; dehydration, clearing and covering, routine. The technic shows particularly the perisomatic fibers, boutons en passant and boutons termineaux. Fibers in nerve tracts may be visible but lightly stained; cell nuclei may be dark, but the cytoplasm remains pale.     

A rapid procedure for routine double staining of cartilage and bone in fetal and adult animals

A simple, rapid procedure for dual staining of cartilage and bone in rodents, particularly in late gestation, has been developed for routine use. The procedure involves rapid, complete skinning of fresh eviscerated specimens following a 30 sec immersion in a 70 C water bath. The unfixed specimen is stained in a mixture of 0.14% Alcian blue and 0.12% alizarin red S in ethanol and glacial acetic acid. Specimens are then macerated in 2% KOH, cleared and hardened in 1:1 glycerin and distilled water, and stored in pure glycerin. Rapid staining of cartilage only is done in a mixture of 0.08% Alcian blue, glacial acetic acid, and ethanol, with subsequent maceration, clearing, and hardening as in the double staining procedure. Rapid staining of bone only, concurrent with maceration of soft tissue, can be done by placing fresh, unskinned specimens in a diluted mixture of alizarin red S in 2% KOH, with subsequent clearing and hardening in 1:1 distilled water and glycerin. Good quality fetal specimens can be prepared for examination by any of these procedures in a minimum of 11/2-2 days as compared to a minimum of 4-5 days for other procedures. Double stained specimens can be examined for abnormalities of the cartilage as well as bone.     

Enhanced Reliability in Silver Impregnation of Terminal Axonal Degeneration-Original Nauta Method

Brains of rat with surgical lesions 3-5 days old are fixed in 10% neutralized formalin (excess of CaCO₃), 20 μ serial frozen sections cut therefrom and kept in neutralized formalin for an additional 24-48 hr. The sections are soaked in distilled water 12-24 hr, transferred to 50% alcohol containing 0.75 ml of concentrated NH₄OH (sp. gr. 0.91) per 100 ml 12-24 hr, placed in distilled water 2-3 hr and then in silver-pyridine solution (AgNO₃ 3% aq., 20 ml; pyridine, 1 ml) for 48 hr. Test sections are transferred directly to each one of 3 ammoniated silver-solutions, pH 12.8, 13.0 and 13.2, made as follows: To 200 ml of solution 1 (silver nitrate, 6.4 gm; alcohol 96%, 220 ml; NH₄OH (sp. gr. 0.91), 28 ml and distilled water, 440 ml) is added respectively 8-12 ml, 12-16 ml and 16-20 ml of solution 2 (2% NaOH) to give the pH desired. The test sections are studied and the optimal ammoniated silver solution chosen. Two baths of ammoniated silver are used, the section placed with continuous agitation into the first bath for 30 sec and the second bath for 60 sec. The sections are then transferred directly into a reducing bath (formalin 10%, 2ml; alcohol 96%, 5 ml; citric acid 1%, 1.5 ml and distilled water, 4.5 ml) for 2 min and from there to 5% Na₂S₂O₃ for 1 min, rinsed in 3 changes of distilled water, dehydrated and mounted.     

A Six-Dye Staining Schedule for Sections of Mesquite and Other Desert Plants

Materials are fixed in FPA (formalin, 2; propionic acid, 1; 70% ethanol, 17). Paraffin sections on slides are brought to 50% ethanol and stained as follows: (1) in Bismarck brown Y, a 0.02% solution in 0.1% aqueous phenol, 10-30 min; wash 30 sec in 0.7% acetic acid, and wash in distilled water 20-30 sec; (2) in crystal violet, 1% in 70% ethanol alkalinized with 1 drop of 1 N NaOH per 100 ml, 12-35 min; wash 30-60 sec in tap water to remove excess stain, and rinse 0.5 sec in 70% ethanol; then mordant in I₂-KI, 1% each in 70% ethanol, 40 sec, and rinse in 70% ethanol 2-5 sec; (3) in a mixture containing 0.4% acid fuchsin and 0.6% crythrosin B in 70% ethanol about 0.5 sec; rinse in 70% ethanol 5-15 sec to remove excess red; dehydrate in 70%, 95%, and absolute ethanol, 2-3 sec each; (4) in fast green FCF, 0.5% in a mixture of equal parts of methyl cellosolve, absolute ethanol, and clove oil, 5-15 sec; rinse in a mixture of clove oil, 10 ml; absolute ethanol, 100 ml; and methyl cellosolve, 10 ml, 5-7 sec; (5) in orange G, 0.75 gm in a mixture of clove oil, 40 ml; absolute ethanol, 40 ml; and methyl cellosolve, 60 ml, 5-30 sec; rinse clean in a 1:1 mixture of xylene and absolute ethanol, 5-20 sec Complete the clearing in pure xylene, 3 changes, 1.5 min in each, and apply a cover glass with synthetic resin. Slides are agitated in all steps except Bismark brown Y, crystal violet, and the xylenes. Contrast and staining intensity are adjusted by varying staining times in the dye solutions.     

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.     

Preparing Specimens of Bone and Teeth for Cutting by the Paraffin Method

A method of preparing bone or teeth for sectioning is described which involves the following steps: 48 hr. in 1:10 formalin; 24 hr. in 70% alcohol; decalcification for several days in 10% HNO₃; rinsing and transferring to 2% potassium alum for 12 hr.; rinsing and treating with 5% NaHCO₃ (or Li₂CO₃) for 24 hr.; washing for 12-24 hr.; then passing through ascending grades of alcohol to xylene. In the case of developing teeth, a slightly different procedure is recommended: fixation in Heidenhain's Susa till hard tissue is decalcified; 24 hr. in 96% alcohol (with three changes); 24 hr. in absolute alcohol (with one change); clearing in xylene or chloroform, and embedding in paraffin.     

Alcoholic Thionin Counterstaining Following Golgi Dichromate-Silver Impregnation

Brains of cats that had been fixed 2 months or longer in 10% formalin were cut into 3-6 mm. slices and impregnated by Golgi's dichromate-silver procedure (6% dichromate solution, 4-6 days; 1.5% silver nitrate solution 2 days). Sections 100 µ thick were cut after embedding in low melting point paraffin. Three changes of xylene and three of absolute alcohol were followed by staining 3-5 minutes in a saturated solution of thionin in absolute alcohol. The sections were dipped quickly in absolute alcohol and cleared in xylene, then differentiation was effected by an equal-parts mixture of absolute alcohol and xylene. A final clearing in three changes of xylene and mounting in Permount completed the process. Counter-staining was most successful when applied to freshly cut sections.     

Basic Two-Dye Stains for Epoxy-Embedded 0.3-1 μ Sections

Dyes used in the 3 methods recommended are: I, thionin and acridine orange (T-AO); II, Janus green and Darrow red (JG-DR); III, methyl green and methyl violet (MG-MV). The first 2 methods were two-solution stains, applied in sequence; the third, required only one solution since methyl violet is present in commercial methyl green. Staining solution and timing was as follows: Method I. 0.1% thionin in a 45% ethanolic solution of 0.01 N NaOH, 5 min at 70 C; rinsing in water and followed by 1 min in a 1% aqueous solution of acridine orange made up in 0.02 N NaOH, also at 70 C, then washed, and dried on slides. Method II. 0.5% Janus green in aqueous 0.05 N NaOH, 5 min at 70 C; rinsing in water then into 0.5% Darrow red in 0.05 N NaOH (aq.), 2 min at 70 C., washing, and drying on slides. Method III. 1% methyl green (commercial, unpurified) in 1% aqueous borax for 15-20 min at 20-25 C, washing and attaching to slides. All staining was performed by floating the sections on the staining solutions, all drying at 70 C, and mounting in a resinous medium. T-AO gave blue to violet cytoplasmic structures, darker nuclei which contrasted strongly with yellow connective tissue and the secretion of goblet cells. JG-DR resembled a hematoxylineosin stain, but by shortening the staining time in DR to 0.5-1 min, collagenous and elastic tissue retained more of the green dye. MG-MV gave dark green nuclei in light green cytoplasm, with collagenous and elastic tissues in blue to violet. As with most methods for staining ultrathin sections, thicknesses of less than 1 μ required longer staining times.     

Modified Clearing Method to Show Sieve Tubes in Minor Veins of Leaves

Leaf samples of Glycine max and numerous other dicotyledonous species were cleared by a common, well established procedure modified by using more concentrated (10% w/v) aqueous NaOH, and by leaving samples in NaOH for 2-4 weeks and in chloral hydrate for 3 days, all at mom temperature. A single dye, chlorazol black E (1g/100 ml absolute ethanol), is used to stain for 3-6 min. Samples are mounted with the lower epidermis upward. Sieve tubes in favorable material can be seen in minor veins and vein endings.     

Preparation of spent brewer's yeast beta-glucans with a potential application as an immunostimulant for black tiger shrimp,Penaeus monodon

Spent brewer's yeast was autolysed and the insoluble yeast cell wall fraction obtained was used as a raw material for the preparation of brewer's yeast beta-glucan (BYG). A simple alkaline extraction was applied and optimized. The BYG with significantly (P相似文献   

Modified clearing method to show sieve tubes in minor veins of leaves

Leaf samples of Glycine max and numerous other dicotyledonous species were cleared by a common, well established procedure modified by using more concentrated (10% w/v) aqueous NaOH, and by leaving samples in NaOH for 2-4 weeks and in chloral hydrate for 3 days, all at room temperature. A single dye, chlorazol black E (1 g/100 ml absolute ethanol), is used to stain for 3-6 min. Samples are mounted with the lower epidermis upward. Sieve tubes in favorable material can be seen in minor veins and vein endings.     

Treatment of molasses wastewater by acetogenic bacteria BP103 in sequencing batch reactor (SBR) system

Acetogenic bacteria BP103 cells could be used as the absorbent for melanoidin pigment (MP) and molasses wastewater (MWW). The maximum MP adsorption yield of this strain observed from the dead (autoclaved) cell. It was two times higher than that with resting cells. However, the MP adsorption yield of the strain was 50-60% decreased by acclimatization with the media containing MP. The deteriorated cells (MP-adsorbed cells) could be recovered by washing with 0.1% SDS, 0.1% Tween 80 and 0.1 mol/L NaOH solutions. Among them, 0.1 mol/L NaOH solution was most suitable according to highest elution ability and no-effect to the MP adsorption capacity (The adsorption yield of deteriorated cell was reduced only 10% after washing three times with 0.1 mol/L NaOH solution). In SBR system, the strain showed very low MP removal yield with both molasses wastewater (MWW) from the anaerobic pond (An-MWW) and stillage from an alcohol factory (U-MWW). However, the MP removal yield was increased by supplementation with carbon sources (glucose). Also, the MP removal efficiency was increased with the increase of supplemented-glucose concentration. The highest COD, BOD(5), TKN and MP removal efficiencies of the SBR system with 10 times-diluted An-MWW solution containing 30 g/L glucose under HRT of seven days were 65.2+/-2.5%, 82.8+/-3.4%, 32.1+/-0.8% and 50.2+/-3.7%, respectively. The large molecular weight fraction of MP in both U-MWW and An-MWW solutions were rapidly removed by acetogenic bacteria BP103, while the small molecular weight fractions of MP still remained in the effluent.     

Injection of Lymphatics: With Colored Cedar Oil; With Plastic

(1) The oil mass consists of: cedar oil, 1; color in oil (a paint pigment, e.g., Prussian blue), 1; and toluene, 2, parts by volume. To use, add 1 ml of diethyl ether to each 10 ml of mass, mix thoroughly and inject into the fresh organ with a very fine glass or metallic needle. Heat the organ in water at 50-60° C before starting the injection, massage gently after injection, then fix. For macroscopic studies, fix 5 days in 5% formalin, and dissect. For microscopic studies, fix at least 5 days in: formalin, 10 ml; Al₂(SO₄)₃, 2 gm; ZnSO₄, 2 gm; acetic acid, 4 ml; and distilled water, 90 ml. Dehydrate with dioxane, embed in paraffin and section at 10-20 μ. Stain with hematoxylin-eosin or with one of the following modifications of Van Gieson's formula: 1. 1% acid fuchsin, 10; picric acid (sat. aq.), 50; and 5% ZnSO₄, 40 volumes. 2. 1% acid fuchsin, 20; picric acid (sat. aq.), 80; and 5% CoSO₄, 40 volumes.

(2) The plastic mass consists of a 5-10% solution of Rhodopas (a vinyl copolymer) in acetone. Injection is made as with the oil mass except that a plastic squeeze-bottle and glass needle is preferable to a syringe. Indirect injection is used for both procedures, i.e., into the organ substance; not into a cannulated lymphatic vessel. After the plastic has hardened (24 hr), the unfixed tissue is subjected to corrosion by 5-10% NaOH in water.     

Ethanol production from cotton-based waste textiles

Ethanol production from cotton linter and waste of blue jeans textiles was investigated. In the best case, alkali pretreatment followed by enzymatic hydrolysis resulted in almost complete conversion of the cotton and jeans to glucose, which was then fermented by Saccharomyces cerevisiae to ethanol. If no pretreatment applied, hydrolyses of the textiles by cellulase and beta-glucosidase for 24 h followed by simultaneous saccharification and fermentation (SSF) in 4 days, resulted in 0.140-0.145 g ethanol/g textiles, which was 25-26% of the corresponding theoretical yield. A pretreatment with concentrated phosphoric acid prior to the hydrolysis improved ethanol production from the textiles up to 66% of the theoretical yield. However, the best results obtained from alkali pretreatment of the materials by NaOH. The alkaline pretreatment of cotton fibers were carried out with 0-20% NaOH at 0 degrees C, 23 degrees C and 100 degrees C, followed by enzymatic hydrolysis up to 4 days. In general, higher concentration of NaOH resulted in a better yield of the hydrolysis, whereas temperature had a reverse effect and better results were obtained at lower temperature. The best conditions for the alkali pretreatment of the cotton were obtained in this study at 12% NaOH and 0 degrees C and 3 h. In this condition, the materials with 3% solid content were enzymatically hydrolyzed at 85.1% of the theoretical yield in 24 h and 99.1% in 4 days. The alkali pretreatment of the waste textiles at these conditions and subsequent SSF resulted in 0.48 g ethanol/g pretreated textiles used.     

Staining of plant Materials Cleared in NaOH

Stains are listed which have proved suitable for staining the epidermis, mesophyll, and sclerenchyma and tracheary elements, respectively, of cleared leaf material of Mouriri and Linociera. Too rapid leaching is avoided by overstaining high in the dehydration series, destaining briefly in the same solvent, and moving through to xylene. Twenty to thirty minutes staining time is generally sufficient. Concentrations and solvents can be varied widely. If destained too much, the material can usually be replaced in the dye with no ill effects. A double stain schedule (Bonnett) of five to ten minutes in 1% Bismarck brown Y in 95% alcohol followed by one to two minutes in 1% fast green FCF in 100% alcohol may be advantageous for thin-walled cells in thick material. It may be preferable to treat thinner material with tannic-acid-iron-chloride followed by safranin (Foster). The effects of bleaches and clearing compounds other than NaOH on staining have not been investigated; however, Dr. Bonnett finds that lactic acid used after NaOH improves clearing and also improves the staining of his combination (above). Mordants can doubtless be used to advantage.     

Ensiled alkali-treated straw. I. Effect of level and type of alkali on the composition and digestibility in vitro of ensiled barley straw

In three experiments barley straw chopped to 5 cm nominal particle length was ensiled in laboratory silos for 90 days after treatment with alkali. In the first two experiments, NaOH was added at 0, 1.05, 2.10, 3.15 or 4.20 g per 100 g straw dry matter (DM) (Experiment 1) or at 0, 2.5, 5.0, 7.5 and 10.0 g per 100 g straw DM (Experiment 2) in solutions at either 60 ml or 120 ml solution per 100 g straw DM. Digestibility in vitro of organic matter (OM) and digestible OM in DM (DOMD) increased with increasing level of NaOH. The effect of volume of solution on digestibility was small. The pH of the straws decreased during storage. The content of neutral detergent fibre decreased as the level of NaOH increased, but there was relatively little change in the contents of acid detergent fibre or acid detergent lignin. Lactate and acetate were detected in all silages, and butyrate was present in silages made from straws treated with less than 5 g NaOH per 100 g straw DM. On opening the silos little moulding was seen and the temperature of the straws remained close to ambient in both experiments throughout 16 days of subsequent exposure to air.In the third experiment, the comparative effects of Ca(OH)₂ and KOH were studied alone and in combination ($\text{50}\text{50}$ by weight) with NaOH. KOH mixed with NaOH gave levels of DOMD in vitro similar to those obtained with NaOH alone. Ca(OH)₂, whilst improving DOMD, was slightly less effective than the other alkalis.The optimum level of alkali for the treatment of barley straw prior to ensiling appeared to be 7.5 g/100 g straw DM. At this level of addition, DOMD in vitro would be expected to be about 65%. Ca(OH)₂ is worth further attention as an alternative to NaOH.     

Clearing, cuticle removal, and staining for the fertile bracts (lemmas and paleas) of grass anthoecia.

The epidermis of the bracts enclosing the flower of grasses contains epidermal cell patterns which are indicative of phylogenetic and systematic relationships among taxa. Treating the heavily cutinized anthoecial bracts (fertile lemma and palea) with 10% NaOH results in the removal of sufficient cuticle to allow examination of the cells of the epidermis. After clearing and removal of the cuticle, the bracts are bleached, washed, dehydrated, and if studied by light microscopy, stained in 2% chlorazol black E and mounted in Diaphane; or, if studied by scanning electron microscopy, dried by the critical-point method and either left uncoated or coated with a film of various conductive metals.     

Clearing, Cuticle Removal, And Staining For The Fertile Bracts (Lemmas And Paleas) Of Grass Anthoecia

The epidermis of the bracts enclosing the flower of grasses contains epidermal cell patterns which are indicative of phylogenetic and systematic relationships among taxa. Treating the heavily cutinized anthoecial bracts (fertile lemma and palea) with 10% NaOH results in the removal of sufficient cuticle to allow examination of the cells of the epidermis. After clearing and removal of the cuticle, the bracts are bleached, washed, dehydrated, and if studied by light microscopy, stained in 2 % chlorazol black E and mounted in Diaphane; or, if studied by scanning electron microscopy, dried by the critical-point method and either left uncoated or coated with a film of various conductive metals.     

Hematoxylin Staining of Tissues Embedded in Epoxy Resins

Sections of 0.5-2 μ thickness are affixed to slides with albumen adhesive, thoroughly dried, and placed in xylene or toluene for 1 hr, then brought through ethanol to water. Sections of tissue fixed in O_sO₄ are treated first in 0.1% KMnO₄, then with 1.0% oxalic acid, and after rinsing, incubated at 60 C for 12-24 hr in hematoxylin (Harris's or Ehrlich's) and counterstained 10-15 min with 0.5% phloxine B. Permanent preparations are made by clearing and mounting in a synthetic resin. The method requires only easily available reagents and is suitable for routine processing of epoxy sections.     

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.