A Hematoxylin and Eosin-Like Stain for Glycol Methacrylate Embedded Tissue Sections

A staining procedure is described for use with glycol methacrylate embedded tissue sections which does not stain the plastic embedment or remove the sections from the glass slides. The basic dye is celestine blue B. It is prepared by treating 1 g of the dye with 0.5 ml concentrated sulfuric acid. It is then dissolved with the following solution. Add 14 ml glycerine to 100 ml 2.5 percent ferric ammonium sulfate and warm the solution to 50 C. Finally adjust the pH to 0.8 to 0.9 The acid staining solution consists of 0.075 percent ponceau de xylidine and 0.025 percent acid fuchsin in 10 percent acetic acid. Slides containing the dried plastic sections are immersed in the celestine blue solution for five minutes and in the ponceau-fuchsin solution for ten minutes with an intervening water rinse. After a final wash, the sections are air dried and coverslipped. This staining procedure colors the tissues nearly the same as hematoxylin and eosin procedures.     

Differentiated Staining of Osmium Tetroxide-Fixed, Vestopal-w-Embedded Tissue in thin Sections

Tissues were fixed at 20° C for 1 hr in 1% OsO₄, buffered at pH 7.4 with veronal-acetate (Palade's fixative), soaked 5 min in the same buffer without OsO₄, then dehydrated in buffer-acetone mixtures of 30, 50, 75 and 90% acetone content, and finally in anhydrous acetone. Infiltration was accomplished through Vestopal-W-acetone mixtures of 1:3, 1:1, 3:1 to undiluted Vestopal. After polymerisation at 60° C for 24 hr, 1-2 μ sections were cut, dried on slides without adhesive, and stained by any of the following methods. (1) Mayer's acid hemalum: Flood the slides with the staining solution and allow to stand at 20°C for 2-3 hr while the water of the solution evaporates; wash in distilled water, 2 min; differentiate in 1% HCl; rinse 1-2 sec in 10% NH,OH. (2) Iron-trioxyhematein (of Hansen): Apply the staining solution as in method 1; wash 3-5 min in 5% acetic acid; restain for 1-12 hr by flooding with a mixture consisting of staining solution, 2 parts, and 1 part of a 1:1 mixture of 2% acetic acid and 2% H₂SO₄ (observe under microscope for staining intensity); wash 2 min in distilled water and 1 hr in tap water. (3) Iron-hematoxylin (Heidenhain): Mordant 6 hr in 2.5% iron-alum solution; wash 1 min in distilled water; stain in 1% or 0.5% ripened hematoxylin for 3-12 br; differentiate 8 min in 2.5%, and 15 min in 1% iron-alum solution; wash 1 hr in tap water. (4) Aceto-carmine (Schneider): Stain 12-24 hr; wash 0.5-1.0 min in distilled water. (5) Picrofuchsin: Stain 24-48 hr in 1% acid fuchsin dissolved in saturated aqueous picric acid; differentiate for only 1-2 sec in 96% ethanol. (6) Modified Giemsa: Mix 640 ml of a solution of 9.08 gm KH₂PO₄ in 1000 ml of distilled water and 360 ml of a solution of 11.88 gm Na₂HPO₄-2H₂O in 1000 ml of distilled water. Soak sections in this buffer, 12 hr. Dissolve 1.0 gm of azur I in 125 ml of boiling distilled water; add 0.5 gm of methylene blue; filter and add hot distilled water until a volume of 250 ml is reached (solution “AM”). Dissolve 1.5 gm of eosin, yellowish, in 250 ml of hot distilled water; filter (solution “E”). Mix 1.5 ml of “AM” in 100 ml of buffer with 3 ml of “E” in 100 ml of buffer. Stain 12-24 hr. Differentiate 3 sec in 25 ml methyl benzoate in 75 ml dioxane; 3 sec in 35 ml methyl benzoate in 65 ml acetone; 3 sec in 30 ml acetone in 70 ml methyl benzoate; and 3 sec in 5 ml acetone in 95 ml methyl benzoate. Dehydrated sections may be covered in a neutral synthetic resin (Caedax was used).     

A Tri-Basic-Dye Stain for Nerve Cells

A new staining method has been developed for the study of nerve cells and Nissl granules which combines three basic dyes, cresylecht violet, toluidine blue and thionin. The use of this tri-basic-dye stain results in finished preparations that are critically stained and permanent. Paraffin sections (4 μ sections preferably) are mounted on slides by the starch medium, deparaffinized and stained by the tribasic staining solution. After differentiation in acidified distilled water, sections are dehydrated, returned to stain solution and again dehydrated, then cleared and mounted in Clarite. Various vertebrate material including normal and pathological human tissues have been stained with this triple dye solution. Especially for pathological material, re-immersion of slides in the staining and 80% alcohol solutions before mounting, differentially intensifies the staining reaction. Fixatives used were 10% formalin, 95% alcohol, Bouin and formalin-Bouin (10% formalin followed by Bouin).     

Techniques for studying adipocytes

Various fixatives as well as tissue and slide handling procedures have been evaluated in attempts to demonstrate adipocytes histochemically while maintaining cell and tissue integrity. The optimal procedure for analysis of immature adipose depots consists of the following steps: 1) fresh, unfixed tissues are rapidly in isopentane quenched in a liquid nitrogen bath; 2) cryostat sections are cut, removed from the knife with a room temperature slide, and then air dried for 5-10 minutes; 3) slides can be stained directly with picro-Ponceau or toluidine blue procedures or with oil red O following fixation for 30 minutes in cold (4 C) 10% formalin-CaCl2 (1.25%). For analysis of mature rat adipose depots steps 2 and 3 are modified as follows: 2) cryostat sections are removed from the knife with a cold slide (-20 C) and dried for 30 minutes at 4 C; 3) the mounted sections are stained with oil red O following fixation for 30 minutes in cold (4 C) 10% formalin-HgCl2 (2.5%). When procedures described above for immature adipose depots are combined with esterase staining, adipocyte cytoplasm is clearly demonstrated. These procedures allow the routine use of fresh frozen, unfixed cryostat sections in studies of adipose cellularity.     

A Tribasic Stain for Thin Sections of Plastic-Embedded, OsO4-Fixed Tissues

Thin (0.5-1 μ) sections of plastic-embedded, OsO₄-fixed tissues were attached to glass slides by heating to 70 C for 1 min. A saturated solution combining toluidine blue and malachite green was prepared in ethanol (8% of each dye) or water (4% of each dye). Methacrylate or epoxy sections were stained in the ethanol solution for 2-5 min. The water solution was more effective for some epoxy sections (10-80 min). Epoxy sections could be mordanted by 2% KMnO₄, in acetone (1 min) before use of the aqueous dye, reducing staining time to 5-10 min and improving contrast. Aqueous basic fuchsin (4%) was used as the counter-stain in all cases; staining time varied from 1-30 min depending upon the embedding medium and desired effects, methacrylate sections requiring the least time. In the completed stain, nuclei were blue to violet; erythrocytes and mitochondria, green; collagen and elastic tissue, magenta; and much and cartilage, bright cherry red. Sections were coated with an acrylic resin spray and examined or photographed with an oil-immersion lens.     

Simplified Aldehyde-Fuchsin Staining of Neurosecretory Cells

Gomori's original aldehyde-fuchsin method has been modified by the combination of Halmi's counter stain with Gabe's preparation, consisting of basic fuchsin, 1 gm; boiling water, 200 ml; with HC1, 2 ml and paraldehyde, 2 ml added after cooling and filtering. The solution so made was allowed to ripen 3-4 days at room temperature, and the precipitate which formed was filtered off and dried at 55-60°C. The staining solution consisted of 0.5 gm of the dry precipitate dissolved in 100 ml of 70% alcohol. The staining follows original procedures except that it is very important to bring slides from water to 70% alcohol before placing them in the aldehyde-fuchsin solution and also to remove all excess staining solution by rinsing in 95% alcohol after staining. The staining solution is stable for at least 6 mo.     

Staining Paraffin Sections Without Prior Removal of the Wax

Paraffin sections are usually rehydrated before staining. It is possible to apply aqueous dye solutions without first removing the wax. Staining then occurs more slowly, and only if the embedding medium has not melted or become unduly soft after catting. To avoid this problem, sections are flattened on water no hotter than 45 C and dried overnight at 40 C. Minor technical modifications to the staining procedures are needed. Mercury deposits are removed by iodine, and a 3% solution of sodium thiosnlfate in 60% ethanol is used to remove the iodine from paraffin sections. At room temperature, progressive staining takes 10-20 tunes longer for sections in paraffin than for hydrated sections; at 45 C, this can be shortened to about three times the regular staining time. After staining, the slides are rinsed in water, air dried, dewaxed with xylene, and coverslipped in the usual way. Nuclear staining in the presence of wax was achieved with toluidine blue, O, alum-hematoxylin and Weigert's iron-hematoxylin. Eosin and van Gieson's picric acid-acid fuchsine were effective anionic counterstains. A one-step trichrome mixture containing 3 anionic dyes and phosphomolybdic acid was unsuitable for sections in wax because it Imparted colors that were nninformative and quite different from those obtained with hydrated sections. Advantages of staining in the presence of wax include economy of solvents, reduced risk of overstaining and strong adhesion of sections to slides.     

A One-Solution Triacid General Stain Which Differentiates Oxygenated from Non-Oxygenated Red Blood Corpuscles

Tissues from representative mammals, amphibia and invertebrates were fixed for 5-24 hr in either an aqueous solution of 8% p-toluene sulfonic acid (PTSA) or in 10% formalin to which 5 gm PTSA/100 ml had been added, and processed through embedding in polyethylene glycol 400 distearate in the usual manner. Sections cut at 4-6 μ were floated on 0.2% gelatin containing 1.25% formalin, and spread and dried on slides at a temperature not exceeding 25 C. Wax was removed with xylene, and the sections brought to water through ethanol as usual. The working staining solution was made from three stock solutions: A. Chlorantine fast blue 2RLL, 0.5%; B. Cibacron turquoise blue G-E, 0.5%; C. Procion red M-P, 0.5%—each of which was dissolved in 98.5 ml of distilled water to which 0.5 ml of glacial acetic acid and 0.5 ml of propylene glycol monophenyl ether (a fungicide) had been added. For use, the three solutions were mixed in the proportions: A, 3; B, 4; and C, 3 volumes. Staining time was uncritical, 10-30 min usually sufficing for 6 μ, sections. The chief feature of the staining is the differentiation of oxygenated and nonoxygenated red blood corpuscles, in reds and blues respectively. Connective tissue stained blue or blue-green and mucin, green. Nuclei and cytoplasm stain according to their condition at the time of fixation. The mixed stain keeps well, remaining active after 2 yr of storage.     

New Uses for Calcium Chloride Solution as a Mounting Medium

Fresh cross sections of stems [Psilotum nudum, Coleus blumei, and Pelargonium peltatum] and roots (Setcreasea purpurea) 120 μm thick were fixed in FPA₅₀ (formalin: propionic acid: 50% ethanol, 5:5:90, v/v) for 24 hr and stored in 70% ethanol. The sections were transferred to water and then to 1% phloroglucin in 20% calcium chloride solution plus either hydrochloric, nitric, or lactic acid in the following ratios of phloroglucin-CaCl₂ solution:acid: 25:4, 20:2, or 15:5. The sections were mounted on slides either in one of the three mixtures or in fresh 20% calcium chloride solution. A rapid reaction of the acid-phloroglucin with lignin produced a deep red color in tracheary elements and an orange-red color in sclerenchyma. Fixed and stored leaf pieces from Nymphaea odorata were autoclaved in lactic acid, washed in two changes of 95% ethanol, transferred to water, and treated with the three acid-phloroglucin-calcium chloride mixtures. The abundant astrosclereids stained an orange-red color similar to that of sclerenchyma in the sections. In addition, a new method is reported for specifically staining lignified tissues. When sections or leaf pieces are stained in aqueous 0.05% toluidine blue O, then placed in 20% calcium chloride solution, all tissues destain except those with lignified or partially lignified cell walls. Thus, toluidine blue O applied as described becomes a reliable specific test for lignin comparable to the acid-phloroglucin test.     

Combined Bright Field and Fluorescence Staining of Paraffin Sections for Studying the Fertilization Process in Plants

PAS-toluidine blue O—aniline blue staining of paraffin sections allows study of histological and cytological detail while retaining aniline blue induced fluorescence in all “callose sites”. Because most autofluorescence is eliminated by the PAS-toluidine blue prestaining, the detail and contrast of the fluorescence image is superior to slides stained in aniline blue alone. Slides are stained by the PAS reaction, 0.03% toluidine blue O, alkaline 0.005% aniline blue, and mounted directly in aqueous mounting medium.     

Serial Sections of Cooked Rice Embedded in Carbowax

Serial sections of cooked rice kernels may be obtained by following either of two dehydration schedules and embedding in Carbowax. In the first schedule the cooked, rinsed and drained kernels are immersed several days in a nonaqueous fixative composed of: isopropyl alcohol, 10 ml; propionic acid, 30 ml; acetone, 10 ml; methylal, 40 ml; dioxane, 30 ml; and propylene glycol, 30 ml (Newcomer's, modified), followed by 7 or 8 days in equal parts of propylene glycol, dioxane and glycerol (changed once), and 4 days on a warming table in the same mixture with 5% Carbowax added. The dehydrated kernels are then infiltrated 4-24 hr with a Carbowax embedding mixture. In the second schedule they are immersed several days in an aqueous solution consisting of: propylene glycol, 12.5 ml; polyethylene glycol 400, 12.5 ml; either with 75 ml of water containing 0.1% thymol, or with a mixture of water, 65 ml; formalin, 10 ml; CaCl₂, 1 gm; and CdCl₂, 1 gm; followed by 3 or 4 days in 50% propylene glycol, and 3 or 4 days on a warming table in 80% propylene glycol with 5% Carbowax added. Infiltration is as above. The composition of the embedding mixture is varied according to the temperature and humidity likely to prevail during sectioning. The texture of the wax may be improved by adding small amounts of gum arabic, spermaceti, and glycerol. Serial sections 3-10 μ thick are placed on clean dry slides, and adhesive dropped at the edges of the ribbon of Carbowax until it is dissolved. The adhesive consists of water-glass (concentrated solution), 1 ml; concentrated ammonia, 1 ml; Carbowax, 5 gm; and water, 98 ml. After the slides are dry they are stored, or immersed 10 min in chloroform, collodionized, and passed to staining solutions. Atmospheric conditions affect not only the Carbowax, but also the response to reagents of cooked rice and of sections.     

Techniques for Studying Adipocytes

Various fixatives as well as tissue and slide handling procedures have been evaluated in attempts to demonstrate adipocytes histochemically while maintaining cell and tissue integrity. The optimal procedure for analysis of immature adipose depots consists of the following steps: 1) fresh, unfixed tissues are frozen rapidly in isopentane quenched in a liquid nitrogen bath; 2) cryostat sections are cut, removed from the knife with a room temperature slide, and then air dried for 5-10 minutes; 3) slides can be stained directly with picro-Ponceau or toluidine blue procedures or with oil red O following fixation for 30 minutes in cold (4 C) 10% formalin-CaCl₂ (1.25%). For analysis of mature rat adipose depots steps 2 and 3 are modified as follows: 2) cryostat sections are removed from the knife with a cold slide (-20 C) and dried for 30 minutes at 4 C; 3) the mounted sections are stained with oil red O following fixation for 30 minutes in cold (4 C) 10% formalin-HgCl₂ (2.5%). When procedures described above for immature adipose depots are combined with esterase fining, adipocyte cytoplasm is clearly demonstrated. These procedures allow the routine use of fresh frozen, unfixed cryostat sections in studies of adipose cellularity.     

Isolation of Avian Trypanosomes by Selective Centrifugation and Subsequent Processing for Electron Microscopy

Paraffin sections are usually rehydrated before staining. It is possible to apply aqueous dye solutions without first removing the wax. Staining then occurs more slowly, and only if the embedding medium has not melted or become unduly soft after catting. To avoid this problem, sections are flattened on water no hotter than 45 C and dried overnight at 40 C. Minor technical modifications to the staining procedures are needed. Mercury deposits are removed by iodine, and a 3% solution of sodium thiosnlfate in 60% ethanol is used to remove the iodine from paraffin sections. At room temperature, progressive staining takes 10–20 tunes longer for sections in paraffin than for hydrated sections; at 45 C, this can be shortened to about three times the regular staining time. After staining, the slides are rinsed in water, air dried, dewaxed with xylene, and coverslipped in the usual way. Nuclear staining in the presence of wax was achieved with toluidine blue, O, alum-hematoxylin and Weigert's iron-hematoxylin. Eosin and van Gieson's picric acid-acid fuchsine were effective anionic counterstains. A one-step trichrome mixture containing 3 anionic dyes and phosphomolybdic acid was unsuitable for sections in wax because it Imparted colors that were nninformative and quite different from those obtained with hydrated sections. Advantages of staining in the presence of wax include economy of solvents, reduced risk of overstaining and strong adhesion of sections to slides.     

Specific Staining of Glycogen with Haematoxylin and Certain Anthraquinone Dyes

Specific staining of glycogen in rat liver fixed in chilled 80% alcohol, chilled formol alcohol or 10% neutral formalin has been accomplished with acid alizarin blue SWR, alizarin brilliant blue BS, alizarin red S, gallein, haematein, and haematoxylin solutions. TO prepare a staining solution, 1 gm dye, 1 gm K₂CO₃ and 5 gm KCl were dissolved by heating in 60 ml of water. Concentrated NH₄OH (0.880 sp.gr.), 15 ml, followed by 15 ml of dry methanol were added to 20 ml of the cooled solution. Paraffi sections were stained for 5 min, rinsed in dry methanol, cleared in xylene, and mounted in D.P.X. The high specificity obviated the need for counterstaining: nuclei and cytoplasm were unstained. Precipitation of stain onto the slide was rare. As all the dyes carried, like carminic acid, numerous groups capable of forming hydrogen bonds, it is suggested that the staining mechanism involved hydrogen bonding.     

Preparing Epoxy Sections of Insect Tissues for Light Microscopy

Sections of aldehyde-fixed and osmium-stained insect tissues embedded in various epoxy resins were affixed to glass slides by use of a slide cover and hotplate combination. A high concentration of solvent vapor over the sections was thus maintained while they dried down on the slides, resulting in excellent flatness and adhesion. Sections were then stained at an elevated temperature with a mixture of equal parts of 3 dye solutions: 1% toluidine blue O, 1% safranin O, and saturated auramine O, all made up in 1% solution of borax in water. The method resulted in excellent differentiation of all insect tissue components including lightly chitinized structures.     

Histochemical Demonstration of Pyrophosphatase

Fresh tissue slices were fixed in 5% formalin containing 0.9% NaCl for 10-20 min and frozen sections therefrom floated for 3 hr at 37°C on an incubating mixture made as follows. Sodium pyrophosphate (Na₄P₂O₇-12H₂O), 1.088 gm was dissolved in 20-30 ml of distilled water and to this was added ferric chloride (FeCl₃-6H₂O), 0.61 gm dissolved in 10-15 ml of water. The precipitate was just dissolved by cautiously adding 5-10% aqueous Na₂CO₃ solution and the pH adjusted to 7.2 with 1N HCl. The volume was made up to 100 ml and 0.9 gm of NaCl added. Before use, 1 ml of 10% Mg(NO₃) was added. After incubation, sections were washed 10-15 min in 0.9% NaCl, then mounted on glass slides and air-dried. When dry, the slides were immersed in 0.9% NaCl containing 0.2-0.5% ammonium sulfide for 2-3 min, then dehydrated rapidly through graded alcohols, cleared, and covered in balsam. Sites of pyrophosphatase activity stained in various shades of green. Acid pyrophosphatase also was histochemically demonstrated by the same principle, excepting that the substrate solution was adjusted to pH 3.7-4.0 with acetate buffer. The pattern of distribution of pyrophosphatase and glycerophosphatase was almost identical.     

Affixing Polyester Wax Sections to Glass Slides by Celloidin and Cellulose Acetate Applied in Sequence

Sections cut from material embedded in polyester wax can be firmly attached as follows: One drop of a 2% solution of celloidin in amyl acetate is smeared on clean slides, and sections taken from the floatation water onto these slides are dried at room temperature. After drying the slides are immersed in a 2% solution of cellulose acetate in acetone for 1 min, transferred directly to absolute ethanol, through 50% ethanol, and into water. Sections affixed by this method and stained by either hematoxylin-eosin or toluidine blue schedules do not loosen and have negligible background staining.     

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.     

A simple method using alizarin red S for the detection of calcium in epoxy resin embedded tissue

This report presents a simple procedure for staining 1-2 microns epoxy plastic sections of cells and mineralizing matrix present in fetal bovine bone tissue cultures. A 0.3% aqueous toluidine blue O solution was used as a cellular stain and was followed with 2% alizarin red S for the detection of calcium at sites of mineralization. Effects of concentration and pH of alizarin red S on the penetration of epon embedded thick sections were investigated. Optimal staining was achieved with a 2% aqueous alizarin red S solution adjusted to a pH of 5.5-6.5. This staining procedure provides unusually clear contrast between mineral and bone cells in plastic sections for light microscopy.     

Oxazine Dyes. I. Celestine Blue B with Iron as a Nuclear Stain

It is suggested that celestine blue B can stain as a colloidal dispersion, the nuclear specificity of which is controlled by the pH. The staining solution is prepared by adding 0.5 ml of concentrated H₂SO₄ to 1 gm of celestine blue B and dissolving the resultant granular mass in 100 ml of 2.5% ferric alum containing 14 ml of glycerol. Sections of amphibian, avian, and mammalian tissue placed for 1 min in this solution and then rinsed in water show as sharp nuclear staining as that usually produced by hematoxylin. A wide variety of fixatives is permissible. Overstaining is not possible within reasonable limits of exposure and no differentiation nor bluing is required. Both the staining solution and stained slides are stable.