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.     

Advantages in the Use of Aldehyde Fuchsin with Van Gieson's Picro-Fuchsin Counterstaining for Differentiating Bone and Cartilage

Specimens of bone were fixed in 10% neutral phosphate-buffered formalin or in Bouin's fluid and decalcified in 10% formic acid buffered with 10% sodium citrate. Materials were embedded in paraffin and 4-5 μ sections attached to slides were oxidized with 0.5% KMnO₄, decolorized in 1% oxalic acid, stained with aldehyde fuchsin, and counter-stained with Van Gieson's picro-fuchsin. Sections were dehydrated, cleared and mounted in a synthetic resin. Microscopically, the differentiation between bone and cartilage was seen as red and purple respectively, with connective tissue red; muscle and erythrocytes, yellow; and elastic fibres purple. The areas occupied by bone, cartilage and erythrocytes could be compared, and also the depth to which cartilage extended into the ossified sites. The advantages of this staining combination are: good contrasts in colour, ease of applying the stain, and virtual self-differentiation of the staining solutions.     

Toluidine Blue Staining of Coccidia in Cultured Animal Cells

Cultured animal cells infected with various species of Eimeria (coccidia) from chickens are washed in Hanks balanced salt solution (HBSS) and fixed in 5% formalin in HBSS. The fixed cultures are washed briefly in distilled water to remove HBSS salts and then dehydrated in a series of mixtures of 40 to 50% ethanol with increasing concentrations of tertiary butanol (TB) and decreasing concentrations of distilled water. Cultures are placed for 1 min in a mixture of 2 parts ethanol :TB (25:75) and 1 part 0.05% toluidine blue O in McIlvaine buffer (pH 6.0), followed by 1 min in 0.05% toluidine blue O in McIlvaine buffer (pH 6.0). The stained cultures are dipped for 1-2 sec in TB, allowed to dry and mounted permanently on slides. Cover-slip cultures fixed and stained by this procedure 8 years ago have not faded or discolored. The alcohol mixtures, formalin in HBSS, stain and buffer can be prepared in large volumes and stored indefinitely. The staining procedure has proven to be rapid and dependable with a variety of cell types in monolayer cultures in research and teaching applications.     

Staining Streptomyces Scabies in Lesions of Common Scab of Potato

Toluidine blue can be used to stain Streptotnyces scabies distinctively in slide cultures or in the lesions of common potato scab. This staining method is based on the metachromaticism of volutin, a constant constituent of the spores and mycelium of S. scabies. Either sections or smears are fixed in FPA (formalin, 5 parts; propionic acid, 7.5; 50% alcohol, 87.5), stained in a 1:100 dilution of saturated aqueous toluidine blue from 20 minutes to 24 hours, dehydrated in an acetone-xylene series and mounted. Cellular constituents of the potato tuber stain blue or are colorless whereas the mycelium of Streptomyces appears as a series of red volutin spheres in the blue stained cytoplasm. The criteria of volutin and cytoplasmic staining along with the 1 µ diameter of the mycelium make it possible to distinguish Streptomyces from the other microorganisms and cells in the lesion region.     

Frozen Section Micro-Incineration

A method for micro-incineration of frozen sections is described. Material containing diffusible or soluble salts is cut on the freezing microtome and the sections are placed into xylol and mounted out of xylol onto Corex D slides previously filmed with glycerin-gelatin medium. Material containing non-diffusible or insoluble salts can be fixed in 10% formalin before sectioning. Sections of the fixed material are dehydrated thru 50, 70, and 95% ethyl alcohol and mounted out of absolute alcohol onto Corex D slides previously fumed with glycerin-gelatin medium. After mounting by either procedure the sections are incinerated in an electric furnace and the temperature of incineration is dependent on the type of tissues to be incinerated and the character of the salts present. The method is time saving and when no fixation is required the whole procedure can be carried out in one hour.     

A Sudan Black B Myelin Stain for Peripheral Nerves

Blocks of fresh issue were fixed 2 or more days in: cobalt sulfate (or nitrate), 1 gm; distilled water, 80 ml; 10% calcium chloride, 10 ml; and formalin, 10 ml. The fixed tissue was washed thoroughly in tap water, embedded in gelatin, frozen sections cut, and mounted on slides with gelatin adhesive. The sections were stained 15-30 min in a saturated, filtered solution of Sudan black B in 70% alcohol, differentiated in 50% alcohol under microscopic observation, and a cover glass applied with glycerol-gelatin. In thick (50-100 μ) sections, myelin stained green to gray-green and this allowed easy differentiation between nerves and other tissue elements.     

A Method for the Preparation of Serial Thick Sections of Plastic-Embedded Plant Tissues

A procedure is described in which thick sections (2-10μ or more) of plastic-embedded plant tissues are mounted in serial order on slides for use in routine light microscopy. Sections are cut with a steel knife on a rotary microtome while the block and blade are bathed with 40% alcohol. The cut sections are placed, in order, in 50% alcohol in the small wells of modified plastic trays where they become flat, pliable and suitable for subsequent handling. Sections remain separate and in correct order in the trays while they are stained, washed, and prepared for final mounting on slides. Mounting involves a simple and rapid procedure of transferring the sections to a slide and heating first on a 70-75 C hot plate (to slowly evaporate the water around the section and to partially affix the section) and then on a 100 C hot plate. This second heating ensures adhesion when xylene-base mounting media, which tend to loosen weakly adhered plastic from the slides, are used. The technique of staining the sections loose provides the following advantages: (1) the problems of section loss and entrapment of stain between section and slide during staining are eliminated, (2) relatively high staining temperature, akalinity, and alcohol concentration of the stain solvent (all of which promote loosening of pm-affixed sections from slides during staining) is allowed, and (3) staining is more even and selective. The procedure has been found to be reliable and fast enough to be of value in a significant variety of routine light microscope studies.     

Cytochemical Staining of Sections from Plastic-Embedded Flagellates

Dinoflagellate chromosomes in sections of plastic-embedded cells were stained without removing the plastic. Azur B and Feulgen procedures were used to localise DNA. Azur B was used with Araldite or methacrylate sections by staining in 0.2% stain in 0.05 M citrate buffer at pH 4 for 1 hr at 50 C followed by rinsing in tertiary butyl alcohol to differentiate the chromosomes. Feulgen stain was used with Araldite sections by hydrolyzing in 1 N HCl at 60 C for 10 min, rinsing in water, staining for 24 hr, washing well, drying and covering. Fast green was used with methacrylate sections to stain proteins by flooding the slide with a 0.1% solution of stain in 0.06 M phosphate buffer at pH 8, allowing the stain to dry out at 40-50 C, washing well, drying and covering. Controls were carried out on material fixed in formalin and treated with nucleases or proteolytic enzymes prior to embedding, and staining.     

A method for the preparation of serial thick sections of plastic-embedded plant tissues.

A procedure is described in which thick sections (2-10 mu or more) of plastic-embedded plant tissues are mounted in serial order on slides for use in routine light microscopy. Sections are cut with a steel knife on a rotary microtome while the block and blade are bathed with 40% alcohol. The cut sections are placed, in order, in 50% alcohol in the small wells of modified plastic trays where they become flat, pliable and suitable for subsequent handling. Sections remain separate and in correct order in the trays while they are stained, washed, and prepared for final mounting on slides. Mounting involves a simple and rapid procedure of transferring the sections to a slide and heating first on a 70-75 C hot plate (to slowly evaporate the water around the section and to partially affix the section) and then on a C hot plate. This second heating ensures adhesion when xylene-base mounting media, which tend to loosen weakly adhered plastic from the slides, are used. The technique of staining the sections loose provides the following advantages: (1) the problems of section loss and entrapment of stain between section and slide during staining are eliminated, (2) relatively high staining temperature, alkalinity, and alcohol concentration of the stain solvent (all of which promote loosening of pre-affixed sections from slides during staining) is allowed, and (3) staining is more even and selective. The procedure has been found to be reliable and fast enough to be of value in a significant variety of routine light microscope studies.     

Zinc Chromate Modification of the Golgi Technic

A modification of the Golgi technic is described in which the reaction takes place in well fixed formalin material. Thin slices (whole sections of adult monkey, cat and rat cerebrum) 2 to 3 mm. thick, from brains fixed 3 to 4 months in 10% formalin, are chromated for two days in 3 g. of zinc chromate dissolved in 98 ml. of distilled water and 2 ml. of formic acid. Slices are then removed, blotted dry and immersed, suspended by a thread, in 0.75% silver nitrate solution for two days. Solution should be changed after the first day. After silvering, the slices are dehydrated rapidly (total time about one hour) in 95% and absolute alcohol, placed in xylene 10 minutes, in low melting point paraffin 10 minutes and embedded in low melting point paraffin. Only surface infiltration is necessary since sections are cut 90 to 100 u. Sections are collected in 95% alcohol, dehydrated in absolute alcohol, cleared in several changes of xylene and mounted in Fisher's Permount. Results with fetal and new born material were not good.     

The Differentiation of Placoid, Ctenoid and Cycloid Scales by Means of Alizarin Red S

This technic has been used successfully by the author for staining, in situ, the placoid scales of selachians and the smaller forms of cycloid and ctenoid scales of teleosts. Sections of skin are dissected from the ventrum of the specimen, cleaned of fascia and muscle tissue and fixed in a 10% formalin solution. The section is macerated in several changes of a 3% KOH solution until translucent. Staining is accomplished in a fresh 2% KOH solution to which is added a saturated alcoholic solution of alizarin red S. The section remains in the stain for 24 hours. If necessary, the tissue may be quickly destained in acid alcohol (1% H₂SO₄ in 95% alcohol). The skin is dehydrated in cellosolve or the alcohol series and cleared in methyl salicylate. Placoid and teleost scales prepared in this manner are stained a brilliant red. The various parts of the denticle are well differentiated.     

Cytochemical Staining of Sections from Plastic-Embedded Flagellates

Dinoflagellate chromosomes in sections of plastic-embedded cells were stained without removing the plastic. Azur B and Feulgen procedures were used to localise DNA. Azur B was used with Araldite or methacrylate sections by staining in 0.2% stain in 0.05 M citrate buffer at pH 4 for 1 hr at 50 C followed by rinsing in tertiary butyl alcohol to differentiate the chromosomes. Feulgen stain was used with Araldite sections by hydrolyzing in 1 N HCl at 60 C for 10 min, rinsing in water, staining for 24 hr, washing well, drying and covering. Fast green was used with methacrylate sections to stain proteins by flooding the slide with a 0.1% solution of stain in 0.06 M phosphate buffer at pH 8, allowing the stain to dry out at 40-50 C, washing well, drying and covering. Controls were carried out on material fixed in formalin and treated with nucleases or proteolytic enzymes prior to embedding, and staining.     

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.     

Staining Methods For Studying Ingredient Distribution In Baked Cakes

A method is described for preparing cake crumb for sectioning and staining. Previous to embedding, the fat was stained and fixed by exposing small blocks of cake to the fumes from a 5%, freshly-prepared, aqueous solution of osmic acid (OsO₄). This was followed by dehydration in ethyl alcohol and tertiary butyl alcohol, removal of air under vacuum and infiltration with paraffin.

Sections were cut 20 and 9Op thick and mounted with water.

Wax was removed by immersion in xylene. The sections were rehydrated in a series of ethyl alcohol dilutions, from concentrated to dilute, then transferred to distilled water.

Protein was then stained pink by immersion of the slides in an acidified 0.04% water solution of eosin Y, or starch was stained blue with a dilute aqueous solution of iodine. Ten grams iodine and 10 g. KI were dissolved in 25 ml. distilled water. This stock solution was diluted for use one to two hundred times.

The relationship between protein and starch was demonstrated by staining the sections with eosin, differentiating in 50% alcohol and staining with iodine.

When slides of cake crumb were prepared in this way, the fat was stained black, the protein bright pink and the starch granules a dark blue.     

Orcein-Hematoxylin in Iodized Ferric Chloride as a Stain for Elastic Fibers, With Metanil Yellow Counterstaining

Autopsy and biopsy specimens of human skin were fixed overnight in alcoholic Bouin's solution, embedded in paraffin, cut at 7 μ, deparaffinized, hydrated to 70% alcohol, and treated as follows—stained 2 hours in a mixture consisting of: 0.2% orcein in 70% alcohol and 1% HC1 (conc.), 125 ml; 5% hematoxylin in absolute alcohol, 40 ml; 6% FeCl₃ in water, 25 ml; and aqueous I₂-KI (1:2:100), 25 ml—rinsed in distilled water until the excess stain was removed—differentiated in 1.2% FeCl₃, 5-15 sec—washed in running water, 5 min—differentiation completed in 0.01% HC1 acid-alcohol, 1 min—a dip in 95% alcohol—distilled water, 2 min—0.25% aqueous metanil yellow, 5-10 sec—a 95% alcohol dip—dehydrated in absolute alcohol, xylene, and mounted in a resinous medium. The technic combines the orcein of Pinkus' stain and the hematoxylin mixture of Verhoeff into a single staining solution and gives sharp and reliable results for both coarse and extremely delicate elastic fibers. These stain purple; nuclei, violet; and background, yellow. The stain allows the use of formalin, Bouin's fluid and Zenker-formol fixation. The results have been consistent in other primates as well as in man.     

A Differential Staining Method for Elastic Fibers, Collagenic Fibers, and Keratin

A method allowing for the differential presentation of elastic fibers, other connective tissue fibers, epithelial and other types of cytoplasm, and keratin is described. The procedure is based on the affinity of orcein for elastic fibers, of anilin blue for collagenic material, and of orange G for keratin. Bouin-fixed, tissue-mat embedded sections are stained in Pinkus' acid orcein for 1 1/2 hours and rinsed in distilled water. The sections are differentiated in 50% alcohol containing 1% hydrochloric acid, washed in tap and then in distilled water. The sections are next transferred for I to 2 minutes to the anilin blue, orange G, phosphomolybdic acid combination known as solution No. 2 of Mallory's connective tissue stain, diluted 1:1 with distilled water. They are then rinsed in distilled water, quickly passed into 95% alcohol, and dehydrated in absolute alcohol containing some orange G, after which they are cleared and mounted. Within less than two hours sections may be stained and mounted with the following results: elastic fibers — red; collagenic fibers — blue; muscle fibers — yellow; keratin — orange.     

Histological Preparation of the Undecalcified Rat Otic Bulla for Mesoscopic Observations

Otic bullas of the rat, obtained by excision and formalin fixed, are successfully embedded in methylmethacrylate by dehydration and subsequent infiltration with plastic under vacuum. Sections 10 μm thick are obtained by cutting the trimmed and sandpapered acrylic blocks on an LKB multirange microtome. The sections are collected on adhesive tape and stained with a Trichrome stain (modified Weigert-van Gieson). Finally, the sections attached to the tape are mounted on microscope slides with glycerin-gelatin and sealed in the same medium. Serial sections are used for three-dimensional graphic reconstruction.     

Differentiation of Myofibrillae, Reticular and Collagenous Fibrils in Vertebrates

A procedure for the differentiation of the mesenchymal derivatives, myofibrillae, reticular and collagenous fibers is presented. Formol-Zenker fixation (5-12 hours) is followed by the washing, iodinization, dehydration and paraffin embedding steps routine for that fixative with the following modifications. Zirkle's butyl alcohol series is used for dehydration and infiltration with paraffin as well as in the alcohol slide series. Embedding paraffin used is Parawax plus 8-10% bayberry wax. Tissue-exposed surface of paraffin block is soaked in water overnight before cutting serial sections at 3-5μ. Sections are mounted using the dilute albumen method, and the slides, thoroughly dried at 37^oC. overnight, are left at 60^o for 10 minutes to melt the paraffin of the sections. Before staining, the sections are given a preliminary treatment with potassium permanganate and oxalic acid. For reticular staining a 10% silver nitrate bath is succeeded by an ammoniacal silver carbonate solution followed by reduction in 1% neutral formalin, toning in gold chloride and fixing in sodium thiosulphate. Myofibrillae, the sacroplasmic limiting membrane and other sarcous elements are stained by Heidenhain's azocarmine solution, adult tissues at room temperature and fetal tissues at 50 ^oC. Differentiation in phosphotungstic acid is followed by the staining of collagenous fibers. For adult tissue, light green SF (C.C.) is used and for fetal tissue, fast green FCF (C.C). A discussion of the preparation of ammoniacal silver solutions is included. Both stock and used solutions of ammoniacal silver have been in use by the author for over a period of two years.     

Calcium in Cells of Fresh Bone Stained with Glyoxal Bis(2-Hydroxyanil)

Fresh ileum of adult rats and vertebrae and calvariae of newborn rats were immersed in a staining solution containing 0.1 gm of glyoxal bis(2-hydroxyanil) (GBHA) per 2 ml of 3.4% NaOH in 75% ethanol, dehydrated in absolute ethanol, cleared in xylene, and embedded in paraffin. Paraffin sections of stained material, 7 μ thick, were affixed to albumenized slides, immersed in 90% ethanol saturated with Na₂CO₄ and KCN to ensure specificity for calcium, rinsed in 95% ethanol, counterstained in 50% ethanol containing 0.1% methylene blue, dehydrated in absolute ethanol, deparaffinized and cleared in xylene, and mounted in neutral synthetic resin. By this procedure, red Ca-GBHA granules were deposited in goblet and Paneth cells, and in the cytoplasm of osteoblasts, osteocytes, chondrocytes, and periosteal cells of developing bones. Calcium in apatite did not stain. In osseous tissues sectioned in a cryostat or processed by the freeze-dry or freeze-substitution method, epiphyseal chondrocyte calcium was removed, and apatite stained so intensely red that it obscured calcium in the bone cells. Failure of control osseous tissues to stain after immersion in a 1% solution of disodium salt of ethylenediaminetetraacetic acid or in the alcoholic, alkaline solvent of the GBHA solution, indicated that the red granules in the cells of developing bone were due to calcium present in the cells in vivo and not due to absorption of GBHA by tissue components other than calcium, or to diffusion of Ca⁺⁺ during staining. Calcium localized in the cytoplasm and processes of the osteogenic cells suggests the need to re-evaluate the role of osteoblasts as depositors of calcium during osteogenesis.     

Block Staining of Botanical Materials

Plant materials, including coleus stem tips, Psilotum stems and onion root tips, were stained in iron-mordanted celestine blue and safranin (Gray and Pickle 1956) prior to embedding and sectioning. Mitotic figures as well as general morphological and anatomical features are adequately stained by this procedure. The plant materials, subdivided so that the largest dimension is about 8-10 mm, are stained for 12-48 hr. The time is dependent upon the tissue and dilution of the stain used. Excess stain is washed out and the tissues are dehydrated, embedded, sectioned and mounted on slides in the usual manner. Following removal of the wax by xylene the sections may be counterstained, or a cover slip may be added immediately.