Carbowax for Embedding and Serial Sectioning of Botanical Material

Plant material infiltrated with gradually increasing concentrations of Carbowax 400, followed by Carbowax 1540 and finally a 19:1 embedding mixture of Carbowax 1540 and 4000 showed minimum shrinkage. Quantitative measurements of shrinkage in tissue of potato tubers gave the following amounts: fixation and washing, about 4%; transfer from water directly to 70% Carbowax 400, 5176; from water through a graded series (5, 10, 15, 20, 30, 40, 50 and 60% Carbowax) to 70%, only 2.5% shrinkage; with an additional 1.5% occurring in transition to the embedding mixture. Dry ribbons are placed on adhesive-coated (gelatin, 5 gm; water, 120 ml; glycerol, 40 ml; phenol, 2 gm) slides in a humidity chamber. In 10-15 min enough moisture is absorbed by the ribbon to float the sections out gently and bring them in contact with the adhesive. Slides are then dried 5-10 min at room temperature. To remove minor wrinkles, the sections are subsequently flooded with water, then redried 12-24 hr; after which, they are ready for staining.     

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.     

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.     

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.     

Methyl Green-Pyronin for Staining Autoradiographs of Hydroxyethyl Methacrylate-Embedded Lymphoid Tissue

Cells in the spleen in DNA-synthesis were labelled with tritiated thymidine. Tissue was fixed for 12 hr in 10% neutral formalin, washed for 4 hr in tap water and dehydrated through 70% and absolute ethanol. The tissue blocks were infiltrated overnight with a mixture consisting of glycol methacrylate, 80 ml; polyethylene glycol 400, 12 ml; and benzoyl peroxide, 0.27 gm. Specimens were cast in BEEM capsules with the final embedding medium consisting of 42 parts of the infiltration medium and 1 part of an acceleration mixture. This mixture consisted of N,N-dimethylaniline, 1 part and polyethylene glycol 400, 15 parts. The blocks hardened in 30 min and were sectioned with an ultramicrotome fitted with glass knives. Sections were coated with Ilford K5 liquid emulsion and exposed for 2 wk. Methyl green-pyronin staining of autoradiographs was carried out at pH 4.1 in acetate buffer containing 0.5% methyl green (Allied Chemicals) and 0.2% pyronin GS (Chroma). Staining was for 30-60 min, after which sections were washed for 1 min in water, blotted, allowed to dry, and mounted in Canada balsm. The procedure resulted in good quality autoradiographs in which the degree of basophilia of labelled cells could be assessed.     

Hydroxyethyl Methacrylate Combined with Polyethylene Glycol 400 and Water; an Embedding Medium for Routine 1-2 Micron Sectioning

Pieces of tissue, with the largest dimension not exceeding 7 mm, are fixed and dehydrated by the procedures of choice. Two stock solutions: A, for infiltration; and B, the accelerator, are used in embedding. Formulas: A, 80 ml of glycol methacrylate (2-hydroxyethyl methacrylate—Rohm and Haas Co., Philadelphia, Pa.) is mixed well with 12 ml of polyethylene glycol (Carbowax) 400 and 8 ml of water; then 0.27 gm of benzoyl peroxide added, heated to dissolve the peroxide, and allowed to cool to room temperature. B, polyethylene 200 or 400, 15 parts, and N,N-dimethylaniline, 1 part, mixed thoroughly. Tissues are first infiltrated completely with solution A, then cast in a mixture consisting of 42 parts of A mixed with 1 part of B. Polymerization occurs in 45 min to 3 hr, depending on the temperature. In a water bath at 20 C, the time required was found to be about 3 hr; at 25 C, 1.5 hr; and at 30 C, 45 min. The plastic block can be trimmed easily, and sections 1-2 μ thick readily cut. Sections can be attached to slides by water flotation, without adhesive, and should be dried at room temperature. Staining with aqueous solutions of basic and acid dyes, without removing the embedding matrix, is sharp and brilliant. When staining of the matrix by basic dyes occurs, this background stain can be completely removed by differentiating in either 2-butoxyethanol, pure ethanol, or a mixture of the two. A number of histochemical reagents have been found compatible with this embedding procedure.     

Selective Staining of Neurosecretory Material in Semithin Epoxy Sections by Gomori's Aldehyde Fuchsin

The epoxy resin was removed from semithin (1 μm) sections by immersing them for 30 sec in sodium methoxide (Mayor et al., J. Biophys. Biochem. Cytol., 9: 909-10, 1961) and then processed as follows: (1) left for 1-3 hr at 60 C in a mixture of formalin, 25 ml; glacial acetic acid, 5 ml; CrO₃, 3 gm; and distilled water, 75 ml: (2) oxidized 10 min in a 1:1:6 v/v mixture of 2.5% KMnO₄, 5% H₂SO₄ and distilled water: (3) bleached in 1% oxalic acid, and (4) stained for 15 min in aldehyde fuchsin, 0.125% in 70% alcohol, or in a 1% aqueous solution of toluidine blue. The neurosecretory material is selectively stained.     

Infiltrating and Embedding Tissues with Mixtures of Polyethylene Glycols and Polyvinylacetate Resins

The use of water-soluble polyethylene glycol polymers (Carbowax, Hydrowax) as embedding media can be extended and facilitated by incorporating a water insoluble polyvinylacetate resin, AYAF (Union Carbide Co.). A combination of 7.5% resin added by heating to a 3:1 mixture of polyethylene glycols 1540 and 4000 gives blocks which may be cut at 2-3 μ. Sections can be floated and properly expanded on an ordinary water bath in a manner which may be impossible with Carbowax alone because of section fragility. This may require judicious adjustment of surface tension by the prior addition of minute quantities of the wax. On water, polyethylene glycol dissolves out of tissues, which remain supported by the resin. After attachment to albumen-coated slides, residual resin may, at option, be removed by a 1-2 min immersion in methyl alcohol without visible impairment of fat content. Abopon is used for mounting. The method appears suitable for the study of intracellular lipids, particularly in tissues which cannot be conveniently handled after Carbowax alone.     

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.     

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).     

Staining Sex Chromatin; Biebrich Scarlet and Fast Green Fcf as a Mixture Versus Their Use in Sequence

Human skin was fixed in Davidson's solution (95% alcohol, 35; formalin, 20; glacial acetic acid, 10; and distilled water, 35—parts by volume) and sections prepared through paraffin embedding in the usual manner. Stock stains were: I(BS)—Biebrich scarlet, 1 gm in 100 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid and 5 ml of glacial acetic acid were added—and II(FG)—fast green, 0.5 gm in 85 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid, 0.3 gm of phosphomolybdic acid, and 15 ml of glacial acetic acid were added. Experimental staining solutions were prepared in the following proportions of stock BS to stock FG—1:1, 2:1, 3:1, 1:2 and 1:3. Sections were brought to 50% alcohol and stained for 15, 20, 25 and 30 min in each of the five BS-FG mixtures, rinsed in 50% alcohol, then dehydrated in 70%, 95%, and absolute alcohol, 2 min each; cleared in xylene, and covered in balsam. The 2:1 (optimum proportion) combination of BS with FG, acting for 20 min, yielded 97% sex chromatin-positive nuclei in female material. If sections were stained in stock solution BS for 2 min, they could be differentiated by a 20 min treatment in the mordanting component of stock FG (without dye) to give a one-color stain. Such stains gave about the same percentage of sex chromatin-positive nuclei as those obtained by the regular two-color procedure. These modifications are simpler, more rapid, and yield results comparable to previously employed techniques.     

Staining Sex Chromatin; Biebrich Scarlet and Fast Green Fcf as a Mixture Versus Their Use in Sequence

Human skin was fixed in Davidson's solution (95% alcohol, 35; formalin, 20; glacial acetic acid, 10; and distilled water, 35—parts by volume) and sections prepared through paraffin embedding in the usual manner. Stock stains were: I(BS)—Biebrich scarlet, 1 gm in 100 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid and 5 ml of glacial acetic acid were added—and II(FG)—fast green, 0.5 gm in 85 ml of 50% alcohol to which 0.3 gm of phosphotungstic acid, 0.3 gm of phosphomolybdic acid, and 15 ml of glacial acetic acid were added. Experimental staining solutions were prepared in the following proportions of stock BS to stock FG—1:1, 2:1, 3:1, 1:2 and 1:3. Sections were brought to 50% alcohol and stained for 15, 20, 25 and 30 min in each of the five BS-FG mixtures, rinsed in 50% alcohol, then dehydrated in 70%, 95%, and absolute alcohol, 2 min each; cleared in xylene, and covered in balsam. The 2:1 (optimum proportion) combination of BS with FG, acting for 20 min, yielded 97% sex chromatin-positive nuclei in female material. If sections were stained in stock solution BS for 2 min, they could be differentiated by a 20 min treatment in the mordanting component of stock FG (without dye) to give a one-color stain. Such stains gave about the same percentage of sex chromatin-positive nuclei as those obtained by the regular two-color procedure. These modifications are simpler, more rapid, and yield results comparable to previously employed techniques.     

Mercuric Bromphenol Blue Staining of Precipitin Lines in Agar

Lines formed by antibody-organ antigen reactions are stained particularly well by a modification utilizing the mercuric bromphenol blue (MBB) mixture of Mazia et al. (Biol. Bull., 104: 57-67, 1953). The agar covered slides are placed overnight in 0.85% NaCI at 4 C, followed by washing for 2 hr in 0.85% NaCI at 25 C. They are then rinsed for 10 min in distilled water, and dried overnight at 37 C. The precipitin lines are fixed by immersing the slides for 25 min in 95% alcohol, followed by 5 min hydration in distilled water. They are stained for 25 min in MBB mixture (HgCI₂, 10 gm; bromphenol blue, 0.1 gm; 95% ethanol, 100 ml). Excess stain is removed by immersing in acidified alcohol (95% ethanol, 98 ml; glacial acetic acid, 2 ml). Finally, the slides are passed through alcohol and xylene, and resin-mounted under coverslips.     

A Plastic Embedding Medium for Thin Sectioning in Light Microscopy

Tissue blocks with surface areas up to 2 cm² can be sectioned at 1 or 2 μ after embedding in a medium consisting of: methyl methacrylate, 27 ml; polyethylene glycol distearate MW 1540, 6 gm; dibutyl phthalate, 4 ml; and Plexiglas molding powder A-100, 9 gm (added last). The methacrylate mixture is polymerized at 50° C by benzoyl peroxide, 0.8 gm/ 100 ml of methacrylate. The polymerized matrix is transparent and the blocks can be cut on a rotary microtome with a steel knife. The plastic can be removed from sections with acetone prior to staining. Artifacts caused by embedding and sectioning are negligible     

Basic Dye Counterstaining of Sections Impregnated by the Golgi-Cox Method

Celloidin blocks of Golgi-Cox impregnated material are cut at 50 μ, the sections collected in 70% alcohol, transferred to a 3:1 mixture of absolute alcohol and chloroform for 2 min, and then stored in xylene or toluene for at least 3 min, or up to 2 wk until processed further. Mounting is done on glass slides which have been coated with fresh egg albumen diluted in 0.2% ammonia water (or a 0.5% solution of dry powdered egg albumen) and then dried at 60°C overnight. For attachment to these coated slides, sections are first soaked for 2-3 min in a freshly prepared mixture of methyl benzoate, 50 ml; benzyl alcohol, 200 ml; chloroform, 150 ml; and then transferred quickly to the slides by means of a brush. After 2-3 min the chloroform evaporates and the celloidin softens. The slides are then immersed in toluene which hardens the celloidin and anchors the sections to the slides. Alcohols of descending concentrations to 40% are followed by alkalinizations, first in: absolute alcohol, 40 ml; strong ammonia water 60 ml, for 2 min, then in: absolute alcohol, 70 ml; strong ammonia water, 30 ml, for 1 hr. Excess alkali is then removed by 70% and 40% alcohol, 2 min each, and a 10 min wash in running tap water. Bleaching in 1% Na₂S₂O₃, for 10 min and washing again in tap water for 10 min completes the process preliminary to staining. The preparations are then stained for 90 min in an aqueous solution of either 0.5% cresylecht violet, neutral red, or Darrow red, buffered at pH 3.6. Dehydration and differentiation in ascending grades of alcohol, clearing with toluene or xylene, and applying a cover glass with a mounting medium having a refractive index of about 1.61 completes the process.     

Improved Preparation of Polyvinyl Alcohol-HgCl2 Fixative Used for Fecal Smears

Rather than dissolve polyvinyl alcohol (PVA) powder in a modified Schaudinn's fixative (containing glycerol) by prolonged heating and stirring, it was found better to coat the particles of PVA with glycerol first. PVA powder (Delkote brand), 5 gm, mixed with 1.5 ml of glycerol and added to 62.5 ml of distilled water, is allowed to soak 3-16 hr in a stoppered flask, then heated to 70-75 C in a water bath (with swirling) for about 10 min or until solution is complete. A separate solution of the other ingredients: 95% ethanol, 31 ml; HgCl₂ 4.5 gm; and glacial acetic acid, 5 ml, is then added to the heated PVA solution, and swirled in the stoppered flask until the mixture becomes clear. This fixative is easy to make, produces little or no sediment, does not thicken with age, and has a long shelf life. It is strongly adhesive and fixes protozoa as well or better than that made by the older method or purchased ready made.     

Embedding Media for 1-2 Micron Sectioning. 2. Hydroxyethyl Methacrylate Combined with 2-Butoxyethanol

A hydroxyethyl methacrylate monomer medium incorporating 2-butoxyethanol requires 2 stock solutions for embedding. Solution A: 80 ml of hydroxyethyl methacrylate (Rohm and Haas Co., Philadelphia, Pa.) is mixed well with 16 ml of 2-butoxyethanol; 0.27 gm of benzoyl peroxide, the catalyst, is added and permitted to dissolve. Heating to 40-50 C may be used to accelerate its solution. Solution B: polyethylene glycol 200 or 400, 15 parts, and N,N-dimethylaniline, 1 part, are mixed thoroughly. Tissues are dehydrated in the customary manner to absolute ethanol or other comparable dehydrant, infiltrated completely with A, then cast in a mixture consisting of 42 parts of A well mixed with 1 part of B. Polymerizaion occurs in 4-7 hr. In a water bath at 20 C the time required was about 7 hr; at 28 C, 4 hr. This medium is based on the author's water-polyethylene glycol-hydroxyethyl methacrylate monomer medium (Stain Techn., 42: 119-23, 1967).     

A Copper Sulfate-Silver Nitrate Method for Nerve Fibers of Planarians

For staining in toto, planarians are fixed in a mixture of 10 ml of commercial formalin, 45 ml of 95% ethanol and 2 ml of glacial acetic acid. After treatment with 70% ethanol 3-10 days, they are washed in distilled water and immersed in 10% CuSO₄. 5H₂O for 3 hr at 50° C, transferred without washing to 1% AgNO₃ for 1.0-1.5 hr at 50° C; and then developed in: 10 ml of 1% pyrogallol, 100 ml of 56% ethanol and 1 ml of 0.2% nitric acid. Gold toning, 5% Na₂S₂O₃ and dehydration follow as usual. For staining sections, material is fixed in the same fixative, embedded in paraffin and sectioned at 10 μ. After bringing sections to water, they are immersed in 20% CuSO₄. 5H₂O for 48 hr at 37° C; then rinsed briefly in distilled water and placed in 7% AgNO₃ for 24 hr at 37° C. They are washed briefly in distilled water and reduced in: hydroquincne, 1 gm; Na₂SO₃, 5 gm and distilled water 100 ml. Gold toning, followed by 5% Na₂S₂O₃ and dehydration completes the process. Any counterstaining may follow.     

A Pollen Grain Squash Technique for Saccharum and Related Genera

Anthers collected between 9 and 10 AM were treated for 1 hr at 26-28 C with a 0.5% solution of colchicine, washed for 2-4 min in water, placed in 0.002 M 8-hydroxyquinoline for 1 hr, washed in water for 10 min and fixed in: methanol, 60 ml; chloroform, 30 ml; distilled water, 20 ml; picric acid, 1 gm and mercuric chloride 1 gm, for 24 hr. After washing they were hydrolysed in 1 N HCl for 15 min at 60 C, stained in leuco basic fuchsin for 30 min, then smeared on a slide in a drop of acetocarmine. The slides were sealed, stored overnight, the paraffin was removed, and the slide passed through a 1:1 mixture of n-butyl alcohol and acetic acid, then through pure n-butyl alcohol and mounted in Canada balsam. The significant features of this procedure are: (1) use of chromosomes in the haploid condition for karyotype analysis, (2) better exaggeration of constrictions for easier interpretation of chromosome types and (3) good spreading in plants with a large chromosome number.     

A Ribonuclease-Gallocyanin Stain for Sexing Skin Biopsies

Skin biopsies for sexing can be fixed best in 10-15% aqueous formalin or this solution saturated with HgCl₂. Bouin's fluid and all chromate mixtures should be avoided. Celloidin-paraffin double embedding is recommended but not essential. Sections are brought to water, mercurial residues removed if necessary, and then washed in distilled water. They are incubated at 37°C in a ribo-nuclease solution: approximately 1 mg of ribonuclease powder (Light's) in 100 ml of glass-distilled water; boiled 3-5 sec after dissolving, and kept in a refrigerator (usable about a week). The sections are rinsed and incubated at 37°C overnight in gallocyanin-chromalum (Einarson, 1951) made as follows: Dissolve 5 gm of chromalum in 100 ml of distilled water, add 0.15 gm of gallocyanin, shake thoroughly, heat slowly and boil 5 min; cool, filter, and wash through the filter with distilled water until the filtrate reaches 100 ml. This solution is usable at once and keeps at least a month. Sections should be dipped in acid alcohol to clean (optional), but no attempt made to differentiate them, and washed in tap water. Dehydration, clearing and covering complete the process. The method is nearly as precise as the Feulgen and more convenient and reliable for routine use on miscellaneous material.