The Use of Picric Acid with the Gram Stain in Plant Cytology

The technic described involves the use of a saturated solution of picric acid in absolute alcohol in the process of dehydration following the gentian-violet-iodine stain as applied to plant cytological material. The method is suitable for both paraffin sections and smears of pollen mother cells fixed in Navashin's or Flemming's solutions. Differentiation in clove oil is very easy since cytoplasm destains immediately, while chromatic material destains very slowly following picric acid. Chromosomes are stained more distinctly than with the usual Gram stain and do not fade.     

Modified Whipf's Polychrome: A Connective Tissue Stain with Special Application for Demonstrating Leishmania

A polychrome stain procedure was developed to demonstrate amastigotes of the protozoan parasite Leishmania braziliensis as well as cytoplasmic and other tissue components in cutaneous lesions of infected animals. The procedure is as follows: stain nuclei for 10 minutes with an iron hematoxylin containing 0.5% hematoxylin and 0.75% ferric ammonium sulfate dissolved in 1:1 0.6 N H₂SO₄:95% ethanol; rinse 4 minutes in distilled water. Cytoplasmic staining is achieved by exposing tissues for 10 minutes to a solution containing 0.25% Biebrich scarlet, 0.45% orange G, 0.5% phosphomolybdic acid and 0.5% phosphotungstic acid in 1% aqueous acetic acid. These first two solutions are modified from Whipf's polychrome stain. Sections are differentiated for 10 seconds in 50% ethanol, rinsed in water, stained 3 minutes in 0.1% aniline blue WS in saturated aqueous picric acid, rinsed in water and differentiated for 1 minute in absolute ethanol containing 0.05% acetic acid. Mordanting overnight in 6% picric acid in 95% ethanol produced optimal results.

This procedure was applied to sectioned material from experimental animals with various protozoa. Trypanosoma cruzi, Besnoitia Jellisoni, Toxoplasma gondii and especially Leishmania braziliensis were well demonstrated. Combining cytoplasmic dyes and phosphomolybdic-phosphotungstic acids into one solution afforded differential staining of tissues by Biebrich scarlet and orange G; connective tissues were stained by this solution. Substantially improved definition of connective tissues resulted after counterstaining. This procedure differs from the Massou sequence in which connective tissues are first stained by cytoplasmic dyes along with other tissues and then destained prior to specific counter-staining. in comparing dyes structurally related to Biebrich scarlet, it was found that Crocein scarlet MOO, but not Poncenu S, was an acceptable substitute. Sirius supra blue GL and Sirius red FSBA were not useful as replacements for aniline blue WS in this procedure.     

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.     

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

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

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

Differential Staining of Two Subpopulations of Purkinje Neurons in Rat Cerebellum with Acid Dyes

We present a new method that stains differently two subpopulations of Purkinje cells in the adult rat. Deparaffinized sections of cerebella, fixed by perfusion with buffered glutaraldehyde or Bouin's fluid were stained with 0.5% light green in 50% ethanolf 10-30 min). The excess dye was removed with saturated aqueous picric acid (10-30 min). At this point some Purkinje cells appeared as lightly stained neurons, while others were strongly stained. Slides were immersed in 0.5% aqueous acid fuchsin for approximately 1 min until the lightly stained neurons acquired a red color. Following immersion in 1% phosphotungstic acid, slides were rapidly dehydrated in ethanol, passed to xylene and mounted in Canada balsam. Two subpopulations of Purkinje cells differing in their protein content in somata and proximal dendrites stained differentially by this method. They occurred in all coronal and sagittal sections and in patches or stripes. Their relative proportion varied from lobule to lobule. A second staining method used potassium permanganate as the sole staining reagent. The staining reagent can be used on sections previously stained with the acid dyes. Purkinje cells appeared as subsets of brownish to deep brown stained neurons, the latter ones corresponding to green stained cells in the dichromic method. The results obtained indicated that the subpopulations reflect real differences among individual neurons and are not artifacts. The technique holds promise for identifying and localizing subsets of Purkinje cells differing in their protein content under normal and experimental conditions and for their further characterization by combined staining and histochemical procedures.     

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.     

Staining Sections Coated with Radiographic Emulsion; A Nuclear Fast Red, Indigo-Carmine Sequence

The radioautographs consisting of emulsion-coated sections, after development, fixation and washing, are stained 3 min (uncritical) by an aqueous solution of 5% aluminium sulfate containing 0.1% nuclear fast red, then washed 2-5 min, and stained 5-10 sec by a saturated aqueous solution of picric acid to which 0.25% of indigo-carmine had been added. This technic stains differentially cell nuclei, cytoplasm, muscle fibers, connective tissues and borders of some epithelial cells. It is unnecessary to subject the slides to differentiating solutions.     

Picric Acid as A Destaining Agent for Iron Alum Hematoxylin

A saturated aqueous solution of picric acid is used to differentiate paraffin sections and smeared pollen-mother-cells stained in Heidenhain's and Delafield's hematoxylins. The method proceeds as usual, except that the iron alum in the destaining process is replaced by picric acid. Mixtures of picro-sulfuric acid and dilute Delafield's hematoxylin and mixtures of picric acid and aqueous hematoxylin have also been tested, of which the latter yields the better result, but is not as good as the other method described here.     

Synthetic Orcein as a Stain for Chick Embryo Cartilage Matrix

Chick embryo tissues fixed in Bouin's fluid, in 10% formol saline or in 10% formol saline with subsequent mordanting in saturated picric acid containing 3% HgCl₂, were examined as 5 μ paraffin sections after staining with 1% synthetic orcein in 80% ethanol containing 1% HCl (conc.). Orcein defined the young elastic fibres formed in the truncus arteriosus, aorta and other large arteries after the 5th day of embryonic development but also reacted with the matrix of cartilage in all parts of the skeleton from the 3rd day onward. It is thought that a glycoprotein or proteoglycan shared by these two tissues could account for their mutual affinity for orcein.     

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.     

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 Durable Nissl Stain for Frozen and Paraffin Sections

Frozen sections, 25-50 /j. thick, of formalin-fixed nervous tissues are mounted following the Albrecht gelatin technic. Paraffin sections, 15 p., are deparaffinized and transferred to absolute ethanol. The slides are then coated with celloidin. Both frozen and paraffin sections subsequently follow the same steps: absolute ethanol-chloroform (equal parts) for at least 20 min, 95% ethanol, 70% ethanol (1-3 min), then rinsed in distilled water. Sections are stained in Cresylechtviolett (Chroma) 0.5% aqueous solution containing 4 drops of glacial acetic acid per 100 ml, rinsed in distilled water, agitated in 70% ethanol until excess stain leaves the slide, and rinsed in 95% ethanol. Sections are then dehydrated in absolute ethanol, followed by butanol, cleared in xylene, and enclosed in permount.     

The effect of fixation on the chemical extraction of glycogen from rat liver

Synopsis Small samples of rat liver, weighing 15 mg or less, were either (a) frozen in liquid nitrogen or (b) fixed at 4°C for 5 min to 2 hr in absolute alcohol, alcoholic picric acid (Rossman's fluid), or aqueous picric acid (Bouin's fluid). The tissue samples were analysed for total glycogen content by a modification of the procedure described by Goodet al. (1933).Comparable yields of glycogen were extracted from freshly frozen and fixed tissue samples. The time of fixation had no apparent effect on the amount of glycogen that could be extracted chemically. Dissolved glycogen was not detectable in the fixatives.It is concluded that (a) the fixatives used in this study do not significantly affect the yield of chemically extractable glycogen from liver; (b) fixation is extremely rapid; and (c) alcoholic fixatives are not significantly superior to aqueous picric acid fixatives for preservation of chemically extractable glycogen in very small samples of tissue.     

A Rapid Combined Fixing and Staining Method for Plant Chromosome Counts

A new method for rapidly preparing slides suitable for chromosome counts by the use of a combined fixing and staining solution involves the substitution of anthraquinone for picric acid in Bouin's formula and the addition of alizarin red S with a metallic salt as a mordant. The fixed smears, after being dehydrated to 95% alcohol, are differentiated in 0.5% sulfuric acid in 95% alcohol saturated with picric acid, washed, cleared and mounted in xylol-balsam. Cymene may be used to intensify the stain. Root tips fixed in the above solution may be dehydrated in dioxan, a paraffin solvent; infiltrated, embedded, sectioned and mounted in the usual way. The sections are subsequently differentiated in picro-sulfuric acid alcohol and cymene. An alternative method of differentiation for this stain is also described.     

Staining Plant Tissues with Chlorazol Black E and Pianese III-B

The staining schedule was developed for a study of the mycorrhizae of red pine, Pinus resinosa Ait. From 70% alcohol, sections are stained in a saturated solution of chlorazol black E in 70% alcohol, 10-30 min; free dye removed by washing in 95% alcohol; stained 18-24 hr in Pianese III-b; rinsed in 95% alcohol, acidified by the addition of 2 ml of saturated aqueous picric acid per 100 ml, 3-4 changes or until the last change is pale yellow or light green; and rinsing in 95% alcohol to remove the acid. If the acid fuchsin is too intense, a cautious differentiation with 95% alcohol containing 1-3% of a 0.1 N solution of NaOH is made. If too much chlorazol black is removed, the effect can be compensated by overstaining with this dye at the beginning of the process. Sections are dehydrated, cleared, and covered in the usual manner. This stain has applications to plant tissues generally, and is particularly effective for meristematic tissues. It shows details of cytoplasmic structures and gives sharp delineation of primary cell walls.     

Counterstaining Golgi-Cox Impregnations with Luxol Fast Blue as a Myelin Stain

Immerse pieces of brain tissue 4 wk in solutions A and B, mixed just before use: A. K₂Cr₂O₇, 1 gm; HgCl₂, 1 gm; boiling distilled water, 85 ml. Boil A for 15 min, cool to 2 C and add: B. K₂CrO₄, 0.8 gm; Na₂WO₄, 0.5 gm; distilled water, 20 ml. Rinse in water and immerse 24 hr in LiOH, 0.5 gm; KNO₃, 15 gm; distilled water, 100 ml. Wash 24 hr in several changes of 0.2% acetic acid and then for 2 hr in tap water. Dehydrate and embed in celloidin. Process a 60 μ section through 70 and 95% ethanol, a 3:1 mixture of absolute ethanol and chloroform, and toluene. Immerse it for 5 min in a solution containing methyl benzoate, 25 ml; benzyl alcohol, 100 ml; chloroform, 75 ml. Orient the section on a chemically clean slide and let air-dry 5-10 min. Process through toluene, 3:1 ethanol-chloroform and 95% ethanol. Place the section for 5-60 min at 60 C in a solution made up of: Luxol fast blue G (Matheson, Coleman and Bell), 1 gm; 95% ethanol, 1000 ml; 10% acetic acid, 5 ml. Hydrate to water and immerse in 0.05% Li₂CO₃ for 3-4 min. Differentiate in 70% ethanol and place in water. Immerse for 5-15 min in a mixture of two solutions: A. cresylechtviolet (Otto C. Watzka, Montreal), 2 gm; 1 M acetic acid, 185 ml; B. 1 M sodium acetate, 15 ml; distilled water, 400 ml; absolute ethanol, 200 ml. Dehydrate to 3:1 ethanol-chloroform. Clear in toluene and apply a coverslip. The technique produces fast Golgi-Cox impregnated neurons against a background of counterstained myelinated fibers. Patterns of the myelinated fibers can be used to localize impregnated neurons.     

Staining Sections of Peripheral Nerves for Axis Cylinders and for Myelin Sheaths

Pieces of mammalian nerves 1 to 2 cm. long were placed under moderate tension and fixed 24–48 hours in: picric acid, saturated aqueous, 90 ml.; formalin, 10 ml.; and trichloracetic acid, 25% aqueous, 2 ml. They were washed in water, cut in two and one end stained with 0.04–0.06% osmic acid solution, while the other was dehydrated, embedded in paraffin, and mounted sections from it stained with protargol. The fixing solution used was selected from a number of combinations of acidified picro-formalin as the one most likely to give satisfactory results when followed by both silver and osmic acid. The use of osmic acid solutions of less than 0.1% concentration avoided the overstaining of myelin sheaths seen frequently when stronger solutions were used with material that had been fixed previously. Protargol, 0.5% solution with fast green FCF added to make 0.05% dye in the final concentration, was used to impregnate sections for axis cylinders. Reduction and toning were done as in Bodian's method.     

Staining Sections of Peripheral Nerves for Axis Cylinders and for Myelin Sheaths

Pieces of mammalian nerves 1 to 2 cm. long were placed under moderate tension and fixed 24-48 hours in: picric acid, saturated aqueous, 90 ml.; formalin, 10 ml.; and trichloracetic acid, 25% aqueous, 2 ml. They were washed in water, cut in two and one end stained with 0.04-0.06% osmic acid solution, while the other was dehydrated, embedded in paraffin, and mounted sections from it stained with protargol. The fixing solution used was selected from a number of combinations of acidified picro-formalin as the one most likely to give satisfactory results when followed by both silver and osmic acid. The use of osmic acid solutions of less than 0.1% concentration avoided the overstaining of myelin sheaths seen frequently when stronger solutions were used with material that had been fixed previously. Protargol, 0.5% solution with fast green FCF added to make 0.05% dye in the final concentration, was used to impregnate sections for axis cylinders. Reduction and toning were done as in Bodian's method.     

Protective Paraffin Infiltration of Soft Tissues in Insects to Facilitate Softening of Hard Exoskeletons

This technique can produce serial sections as thin as 5 μ from hard chitin-covered materials of insects or other arthropods. Procedures: Fix with alcoholic Bouin's fluid for 3 hr. Henceforth subject material to partial vacuum in each step to ensure a final proper embedding. Wash with 80% ethanol 2 or 3 times for 2 hr or until the picric acid is largely removed. Dehydrate to 90% ethanol and give 2 changes of n-butanol 2 hr each, and one of a 1:1 n-butanol-paraffin mixture in 56-57° oven for 12 hr. Finally, use 2 baths of pure paraffin, 3 hr each, to complete the infiltration. After the last bath, withdraw the specimen from the paraffin, and remove the superficial paraffin, first mechanically and then with a xylene bath for 4 min. Rinse first with n-butanol, and afterwards with absolute ethanol, 2 min each. The compound eyes are protected with a paraffin covering, the specimen is hydrated with a 1% aqueous solution of detergent for 1 hr and then washed with running tap water. The material is treated with a concentrated sulfuric-nitric mixture (H₂SO₄:HNO₃) for 4 hr to eliminate the exoskeleton. After this treatment, the specimen is washed with running tap water for 12 hr, dehydrated with acetone and then bathed in a 2% solution of celloidin in ethyl acetate to form a protective artificial cuticle. This coating is hardened with 2 quick baths of chloroform, the specimen reembedded in paraffin, and the block cast for sectioning.     

Effect of phosphomolybdic acid on the binding of Sudan black B

Paraffin sections of tissues fixed in absolute alcohol or Carnoy's fluid were mordanted in a 1% aqueous solution of phosphomolybdic acid, stained in saturated solutions of Sudan black B, acetylated Sudan black, various solvent and basic dyes in 70% ethyl alcohol for 5 min at room temperature, dehydrated in alcohol and covered in Permount. Sudan black B and other dyes with basic groups stained basement membranes, reticulum and collagen fibers intensely. Acetylated Sudan black, Sudan IV and oil red 0 did not color any tissue structures. Control sections, without pretreatment, did not bind Sudan black B. These findings indicate interaction between basic groups of the dye and free acid groups of phosphomolybdic acid.