A Simple Histochemical Reaction for Aldehydes

A study has been made on the possibility of replacing leucofuchsin by colored basic fuchsin for the histochemical demonstration of aldehydes. Several tissues from mammals and various pertinent fixatives were used. Aldehydes were freed from carbohydrates by oxidation and from thymonucleic acid by hydrolysis.

It was found that the colored form and not necessarily the leucoform of basic fuchsin can be used histochemically in demonstrating aldehydes. The technic used is as follows: (1) Treat with 1.0-0.5% H₅IO₆ (or in 1% KIO₄ in M/1 H₂SO₄) for 5 to 10 min. and wash thoroughly. For thymonucleic acid hydrolize with N HCl 5 min. at room temperature, 10 min. at 60°C. and 5 min. at room temperature. (2) Stain for 2-3 min. with 0.05% basic fuchsin in 5% ethanol, 3% phenol. (3). Transfer immediately to 1 or 2 changes of 1% sodium bisulphite or potassium metabisulphite in 0.1-0.2 N H₂SO₄ for a total of 5 min. (4) Rinse with water and treat with M H₂SO₄ in 95% ethanol for 3-5 min. 6. Wash thoroughly in water and dehydrate, clear, and mount. For glycogen and mucin the following counterstaining solution is recommended: orange G, 0.25 g.; light green SFY, 0.10 g.; phosphotungstic acid 0.50 g.; 50% ethanol, 100 ml.; glacial acetic acid, 0.25 ml.     

A Consistent Phosphotungstic Acid Hematoxylin Stain for Glial Fibers

After deceration, celloidinization and hydration, oxidize 10 micron paraffin sections for 15 min in a solution containing 0.3 g KMnO₄, and 0.1 ml conc. H₂SO₂, per 100 ml distilled water. Wash in water and reduce in 5% oxalic acid until the sections are colorless. Wash thoroughly in water and place in 4% iron alum solution for two hours. Wash briefly in water and stain for two hours in phosphotungstic acid hematoxylin. Rinse briefly in 95% ethanol and dehydrate in n-butyl alcohol or absolute ethanol for 4 min with two changes, clear and mount. Glial fibers, myofibrils, red blood cells, etc. are stained blue while astrocyte cell bodies, collagen, etc. are stained red. This stain has proven highly consistent in a wide variety of astrocytic derangements. Despite the intensity of this PTAH modification, false positive staining was not observed.     

A Staining Procedure for Use in the Brucella Opsonocytophagic Test

The following staining procedure is recommended for use in the Brucella opsonocytophagic test in order to avoid confusing results obtained with stains of the Hasting or Wright type: Fix spreads for 5 minutes or longer in absolute methyl alcohol. Stain for 10 to 30 minutes in a solution of the following: 0.5 g. NaCl, 0.5 g. phenol, 0.5 g. methylene blue, 0.02 g. Na₂HPO₄+12H₂O, 50 cc. distilled water, 50 cc. methyl alcohol. Wash slides gently in water. Air dry. By this procedure, the bacteria and the nuclei of the leucocytes appear deep blue. The cytoplasm of the leucocytes appears faintly green with the cell outline distinctly visible. Cytoplasmic granules do not stain.     

A Protargol Method for Staining Nerve Fibers in Frozen or Celloidin Sections

Extensive experimentation with protargol staining of neurons in celloidin and frozen sections of organs has resulted in the following technic: Fix tissue in 10% aqueous formalin. Cut celloidin sections IS to 25 μ, frozen sections 25 to 40 μ. Place sections for 24 hours in 50% alcohol to which 1% by volume of NH₄OH has been added. Transfer the sections directly into a 1% aqueous solution of protargol, containing 0.2 to 0.3 g. of electrolytic copper foil which has been coated with a 0.5% solution of celloidin, and allow to stand for 6 to 8 hours at 37° C. Caution: In this and the succeeding step the sections must not be allowed to come in contact with the copper. From aqueous protargol, place the sections for 24 to 48 hours at 37° C. directly into a pyridinated solution of alcoholic protargol (1.0% aqueous solution protargol, 50 ml.; 95% alcohol, 50 ml.; pyridine, 0.5 to 2.0 ml.), containing 0.2 to 0.3 g. of coated copper. Rinse briefly in 50% alcohol and reduce 10 min. in an alkaline hydroquinone reducer (H₃BO₃, 1.4 g.; Na₂SO₃, anhydrous, 2.0 g.; hydroquinone, 0.3 g.; distilled water, 85 cc; acetone, 15 ml.). Wash thoroly in water and tone for 10 min. in 0.2% aqueous gold chloride, acidified with acetic acid. Wash in distilled water and reduce for 1 to 3 min. in 2% aqueous oxalic acid. Quickly rinse in distilled water and treat the sections 3 to 5 min. with 5% aqueous Na₂S₂O₃+5H₂O. Wash in water and stain overnight in Einarson's gallocyanin. Wash thoroly in water and place in 5% aqueous phosphotungstic acid for 30 min. From phosphotungstic acid transfer directly to a dilution (stock solution, 20 ml.; distilled water, 30 ml.) of the following stock staining solution: anilin blue, 0.01 g.; fast green FCF, 0.5 g.; orange G, 2.0 g.; distilled water, 92.0 ml.; glacial acetic acid, 8 ml.) and stain for 1 hour. Differentiate with 70% and 95% alcohol; pass the sections thru butyl alcohol and cedar oil; mount.     

Method for Staining Bacterial Flagella

The following method of staining bacterial flagella is ecommended for use on smears made from suspensions of 10 to 16-tour agar slant cultures, incubated 30 minutes at 37°C before spreadng on thoroly cleaned and named slides:

1. Cover with fixative (100 cc. of 1/4 sat. aqu. solution picric acid, with 5 g. tannic acid and 7.5 g. ferrous sulfate).
2. Wash with tap water, dry and cover with Fontana spirochaete stain; heat to steaming and allow to act for 1 to 2 minutes. Wash in ap water. The stain is prepared as follows: To 25 cc. 2% AgNO₃ add dilute ammonia till the precipitate which forms redissolves; then add more AgNO₃ till a faint turbidity results. A clear solution is useess.

     

Staining Paraffin Sections with Protargol 3. The Optimum pH for Reduction. 4. A Two-Hour Staining Method

Further work on conditions affecting the reduction of paraffin sections impregnated with protargol showed that the optimum pH for sulfite-amidol mixtures was between 6.5 and 7.5. A staining method which requires about two hours to complete consists of the following steps: (1) One hour impregnation at 60° C. in 10% AgNO₃. (2) Wash in distilled water 3 changes of 30 sec. each. (3) Put into protargol (Winthrop Chem. Co., New York, N. Y.) 0.2% aq. for another hour at room temperature. (4) Rinse 2 sec. (5) Reduce one to two min. in amidol 0.2 g., Na₂SO₃ 8 g., NaHSO₃ I g., and water 100 cc. (6) Wash thoroly. (7) Tone with 0.1% gold chloride. (8) Wash. (9) Reduce with a 0.5% aq. soln. of amidol (no sulfite). (10) Wash, dehydrate and cover. The method stains neurofibrillae and unmyelinated fibers and has worked well on most tissues of vertebrates. The stain follows acid alcoholic fixation.     

Mordant Blue 3: a Readily Availablx Substitute for Hematoxylin in the Routine Hematoxylin and Eosin Stain

Mordant blue 3 may be used as a suhstitute for hematoxylin in hematoxylin and eosin stains. The staining solution consists of 0.25 g dye, 40 ml of 10% iron dam, 5 ml of cone H₂SO₄, and 955 ml of dirtilled H₂O. Staining the is 5 minutes, followed by differentiation in acid water or acid alcohol. After blueing, the seaions are counterstained with emin. Results closely resemble the hematoxylin and eosin stain.     

Staining Nuclei and Chromosomes in Protozoa

Technics for free-living forms such as Paramecium and for parasitic forms such as the opalinid ciliates are described.

Paramecium: Fix paramecia in hot Schaudinn's fluid containing 5% of glacial acetic acid for 5-15 minutes. (A hot water bath for maintaining the proper temperature of the fixative is described.) Dehydrate up to 83% alcohol. Mount the specimens on albuminized cover glasses. (A table for mounting animals on cover glasses is described.) Apply a thin layer of collodion to the cover glass to prevent the loss of the specimens during the subsequent handling. Pass through descending grades of alcohol to water. Mordant in 4% iron alum for 24 hours. Stain in 0.5% hematoxylin for 24 hours. Destain in saturated aqueous picric acid. Rinse in tap water, expose to ammonia vapor for a second, and then rinse again in tap water. Wash in running water for 1 hour. Dehydrate. Clear, then mount in damar.

Opalinid Ciliates: Make smears on cover glasses and fix them while wet. If the opalinids are to be subsequently stained in hematoxylin, fix in hot Schaudinn's fluid (containing 5% of glacial acetic acid) for 5-15 minutes. Pass through descending grades of alcohol to water. Mordant in iron alum for 24 hours. Stain in hematoxylin for 24 hours. Destain in saturated aqueous picric acid. For Feulgen reaction, fix in a modified weak Flemming's fluid for 1 hour. Wash in running water for 30 minutes. Hydrolyze. Leave 3 hours in fuchsin decolorized with H₂SO₃ (Feulgen formula). Wash in H₂SO₃, then in running water for 15 minutes. Dehydrate up to 95% alcohol. Counterstain with fast green FCF for 2 minutes. Dehydrate in absolute alcohol. Clear, then mount in damar.     

A Controllable Silver Stain for Nerve Fibers and Nerve Endings

A paraffin section method is described with a yellow-brown-black color range comparable to that of Ranson's pyridine silver block stain. After impregnation with activated protargol and reduction with a fine grain photographic developer, silver nitrate impregnation and reduction are repeated as often as necessary. The procedure is as follows:

Place hydrated sections of tissue fixed in chloral hydrate (25 g. in 100 ml. of 50% alcohol) in 1% aqueous protargol (Winthrop Chemical Co.) containing 5-6 g. metallic copper for 12-24 hours. After rinsing in 2 changes of distilled water, reduce 5 to 10 minutes in: Elon (Eastman Kodak Co.) 0.2 g., Na₂SO₃, dessicated, 10 g., hydroquinone 0.5 g., sodium borate powder 0.1 g., distilled water 100 ml. Wash thoroly in 4 or 5 changes of distilled water and place in 1% aqueous AgNO₃ for 10-20 minutes at 28°-50° C. Rinse in 2 or 3 changes of distilled water and reduce in the elon-hydroquinone solution. After thoroly washing in 4 or 5 changes of distilled water, examine under microscope.

If too pale, treat again in silver nitrate for 10-20 minutes, rinse, reduce 5-10 minutes and wash thoroly until nerve fibers show distinct microscopic differentiation, then dehydrate, clear and mount.     

Staining of Negri Bodies

The following procedure for staining Negri bodies in sections is based on methods previously described by MacNeal, by Haynes, and by Richter:

Fixation:

1. 1. Zenker's solution 4 hours at 37°C or Dominici's 3 hours.
2. 2. 70% alcohol, 12 to 18 hours at room temperature.
3. 3. 80% alcohol, about 5 to 6 hours.
4. 4. 90% alcohol, about 4 to 6 hours.
5. 5. Absolute alcohol about 16 hours.
6. 6. Ether and absolute alcohol aa, about 8 hours.
7. 7. 16 to 24 hours in the following mixture: celloidin 1 g., methyl salycilate 25 cc., abs. alcohol 25 cc., ether 25 cc.
8. 8. Chloroform and paraffin, 2 to 3 hours.
9. 10. Paraffin, 1 to 1 1/2 hours.
10. 11. Embed.

staining:

1. 1. Cut sections 4 to 5 μ.
2. 2. Bring section to water and cover with Lugol's iodine for 10 minutes.
3. 3. Decolorize with a 2% sodium thiosulfate (hypo).
4. 4. Wash thoroly with water.
5. 5. Cover with a mixture of equal parts of 0.5% phloxine and 1% eosin Y (National Aniline brand) and leave for 15 minutes.
6. 6. Wash with water and stain 2 to 5 minutes in 0.1% azure B (National Aniline).
7. 7. Wash with 96% alcohol and decolorize in a mixture of 2 parts absolute alcohol with 1 part clove oil, ordinarily for not more than 1/2 to 1 minute.
8. 8. Dehydrate rapidly, clear, and mount in Yucatan Elemi.

     

A New Method of Silver Impregnation for Nerve Cells and Fibers

Fragments of tissue, immediately after death, are fixed in Debaisieux's modification of the Duboscq-Brazil picro-aceticformol fluid, and treated as follows: Hydrate by soaking 2-6 hr. in distilled water with 30 drops of cone. NH₄OH per 100 cc. Freeze and cut sections about 25μ in thickness. Bleach sections about 15 min. in ammoniacal water (52 drops cone. NH₄OH per 100 cc. water). Transfer to 20% AgNO₃ solution and heat at 45° C. till light brown. Add cone. NH₄OH drop by drop till the Ag precipitates and then redisolves into an opalescent solution. Pour solution and sections into a little distilled water and transfer sections quickly to formaldehyde solution (3 cc. formalin to 100 cc. water). Dip sections in distilled water and transfer to 1% aqueous gold chloride till deep blue. Place for about 10 minutes in 5% aqueous sodium thiosulfate solution for fixing and clearing. Wash thoroly in tap water, dehydrate and mount. Special directions are given for applying this technic to delicate material such as insects, and for use with serial sections.     

Acid Fuchsin as a Connective Tissue Stain After Phosphomolybdotungstic Mordanting

Certain acid fuchsias stain connective tissue deep red after phosphomolybdotungstic mordanting in a modified Masson procedure, others are entirely unsatisfactory for mis purpose. Spectrophotometric examination gives no reliable criteria for separation of acid fuchsins satisfactory for this purpose from unsatisfactory ones. Sulphonation of basic fuchsin with 3.5 to 4 parts of 25-30% fuming H₂SO₄ to 1 part of dye gives a satisfactory product at temperatures as low as 65 to 70°C. in 30 minutes, while use of 5 to 7.5 parts of acid at this and at higher and lower temperatures gives unsatisfactory products. Satisfactory products may be produced with 15% fuming H₂SO₄ in similar quantities, and even with concentrated H₂SO₄, but some unconverted basic fuchsin remains with both and, with the latter, lower quantities give unsatisfactory products. Brief chemical studies indicate that oversulphonation may occur in the manufacture of acid fuchsin and that this is just as deleterious as undersulphonation.     

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.     

An Evaluation and Modification of Cole's Hematoxylin

Consistency in staining with an alum hematoxylin is possible by the routine use of fresh staining solutions. A modification of Cole's hematoxylin is so easily prepared that fresh staining solutions present no problem. The staining solution consists of 100 ml 1.2% aqueous KA1(SO₄)₂ .12 H₂O, 1 ml 10% alcoholic hematoxylin and 2 ml 1% iodine. Mix, place in paraffin oven overnight and stain sections 5 minutes. The three solutions can be kept as stock solutions for years.     

Staining Reticulin with Gold

This bromine-iodine-gold chloride-reduction sequence stains reticulin in formalin-fixed paraffin sections without risk of sections becoming detached. After hydration, sections are exposed to 0.2% bromine water containing 0.01% KBr for 1 hr, then rinsed and placed for 5 min in a solution consisting of KI, 2 gm; iodine crystals, 1 gm; and distilled water, 100 ml. After this the sections are well washed in distilled water, immersed for 5 min in 1% w/v aqueous solution of chloro-auric acid, again rinsed in distilled water, and the gold is reduced by placing in freshly made 3% H₂O₂ for 2-4 hr at 37 C, or in 2% oxalic acid for 1-3 hr at the same temperature.     

Gross Demonstration of the Mammalian Atrioventricular Bundle by A Periodic Acid-Schiff Procedure

The following procedure stains the atrioventricular conduction system selectively. (1) Wash the fresh heart with physiological saline solution to free it of blood; (2) fix it in 10% formalin containing 0.5% HIO₄ for 1 hr; (3) wash in 3 changes of distilled water for 20 min; (4) keep in 80% alcohol for 12 hr to 2 wk; (5) wash with distilled water; (6) treat with a dilute Schiff's reagent containing 0.1 gm of basic fuchsin per 100 ml for 0.5-2 min; (7) rinse in three changes of 2% Na₂SO₃ in 0.2 N HCI for 3-5 min; (8) wash and examine in 80% alcohol; store in 80% alcohol.     

The Gallocyanin-Chrome Alum Stain; Influence of Methods of Preparation on Its Activity and Separation of Active Staining Compound

A dye, which is probably a cationic chelate, has been separated from a gallocyanin-chrome alum staining solution and prepared in the dry form. This dye is apparently the major staining compound. To prepare the chelate or dye, dissolve 150 mg of gallocyanin and 15 gm of chrome alum in 100 ml of distilled water and boil for 10-20 min, cool, filter, wash the precipitate with sufficient distilled water to restore the volume of the filtrate to 100 ml, then add concentrated NH₄OH until the pH is raised to 8-8.5. Filter, with suction, through a medium porosity fritted glass funnel. Wash with 100-200 ml of anhydrous ethyl ether and air dry the precipitate. This ratio of chrome alum to gallocyanin and the 10-20 min boiling time are optimal for preparation of the staining solution, which may be used either for staining or for separation of the chelate in its dry form. From the dried chelate, the staining solution is prepared as a 3% solution in1 N H₂SO₄ and a staining time of 16-24 hr is required. No differentiation is needed; the stain is self-limiting.     

Improving the Staining Qualities of Toluidin Blue

To a 1.0% filtered aqueous solution of toluidin blue add drop by drop 4-5 ml of either a saturated aqueous solution of HgCl₂ or of KI. Collect the resulting dark precipitate on a filter paper and wash it with numerous small quantities of distilled water applied to both inside and outside of the filter paper. Wash until the drippings are distinctly blue (equivalent to about a 0.05% dye solution). Remove the paper and its contents from the funnel and dry either at room temperature or at 37°C. When dry, the treated dye can be brushed off the paper and stored. To prepare a staining solution add a weighed amount (0.12 gm if derived from the HgCl₂ treatment, or 0.3 gm if from KI) to 100 ml of distilled water. This insures a saturated solution in either case and gives a satisfactory stain with most sections in 10-30 min. Thionin and other members of the thiazine dyes also showed improvement in staining qualities after this treatment.     

Method for Staining Bacterial Flagella

The following method of staining bacterial flagella is ecommended for use on smears made from suspensions of 10 to 16-tour agar slant cultures, incubated 30 minutes at 37°C before spreadng on thoroly cleaned and named slides:

Cover with fixative (100 cc. of 1/4 sat. aqu. solution picric acid, with 5 g. tannic acid and 7.5 g. ferrous sulfate).

Wash with tap water, dry and cover with Fontana spirochaete stain; heat to steaming and allow to act for 1 to 2 minutes. Wash in ap water. The stain is prepared as follows: To 25 cc. 2% AgNO₃ add dilute ammonia till the precipitate which forms redissolves; then add more AgNO₃ till a faint turbidity results. A clear solution is useess.     

凋落物去除和氮添加对亚热带阔叶林土壤不同组分碳、氮的影响

人类活动显著增加了氮沉降,对森林生态系统产生了不同程度的影响;凋落物在其分解过程中输入的大量有机碳、氮也会影响土壤碳氮的形成、稳定及转化.本研究选择亚热带常绿阔叶林,对样地进行8年氮添加[对照(0)、低氮(75 kg·hm^-2·a^-1)、高氮(150 kg·hm^-2·a^-1)]和控制凋落物处理(保留凋落物、去除凋落物),之后采集土壤样品,通过K₂SO₄、Na₂B₄O₇、Na₄P₂O₇、NaOH、H₂SO₄、Na₂S₂O₄、HF等化学试剂逐级浸提土壤,测定各浸提液和残渣中的碳、氮含量,研究凋落物及氮添加对土壤矿物结合态碳、氮的影响.结果表明: 整体上,胡敏素(humin,H)组分的土壤碳、氮含量均为最高,分别占土壤全量的33.5%和33.3%.Na₂B₄O₇溶液提取的土壤可溶性碳、氮含量最高,其次是NaOH和Na₄P₂O₇溶液,3种试剂提取的土壤可溶性总碳、可溶性总氮以及可溶性有机氮分别占提取总量的46.2%、47.9%和76.5%.与对照相比,氮添加增加了Na₂S₂O₄和H组分碳、氮含量;与保留凋落物比较,去除凋落物降低了Na₂B₄O₇、H₂SO₄、Na₂S₂O₄和H组分的碳含量,以及NaOH、HF和H组分的氮含量.保留凋落物和氮添加显著增加了K₂SO₄组分氮含量.可见,保留凋落物和外源氮通过影响化学稳定性不同的土壤组分的碳氮变化来改变土壤碳氮过程.