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.     

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.     

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.     

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.     

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     

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.     

Rapid Single-Solution Polychrome Staining of Semithin Epoxy Sections using Polyethylene Glycol 200 (PEG 200) as a Stain Solvent

Rapid, onestep polychromatic staining of 0.75-1.5 μm epoxy sections of glutaraldehyde-osmium fixed tissues can be obtained with mixtures of basic fucbsin and toluidme blue O in alkaline polyethylene glycol ZOO (PEG ZOO). Sections are attached to slides by heating at 100 C for 45 seconds and stained at that temperature for 2-3 minutes with a solution consisting of PEG 200 (50 ml), 0.2 N KOH (0.75 ml), basic fuchsin (1.7 gm), and toluidine blue O (0.3 gm). Red-blue balance and selective staining of different structures can be controlled by varying the amount of toluidine blue added. After rinsing with 10% acetone and rapid drying, sections are covered with immersion oil or mounting medium and a cover-slip. Total time from cutting of a section to finished preparation is less than 6 minutes. This staining solution is stable, does not produce precipitates on the sections, and does not wrinkle or lift the sections from the slides.     

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.     

Polyethylene Glycol Distearate 600 With 10% 1-Hexadecanol; A Superior Embedding Wax for WARM Climates

A mixture of 90% polyethylene glycol distearate 600 and 10% 1-hexadecanol has a melting point of approximately 43 C and is suitable as a replacement for conventional polyester wax where laboratory temperatures are above 24 C. Sections are attached to slides by floating out on 10% formalin and picked up on slides precoated with undiluted egg albumen. Since the wax is opaque, very small specimens can be embedded after fixation in a block of 0.5% agar before dehydration and embedding in wax, thus facilitating their handling and orientation. The wax mixture sections well with a razor blade in a holder.     

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.     

The Mounting of Carbowax Sections with Lighter-Fluid and Rubber Cement

Serial sections cut from plant tissues embedded in Carbowax have been affixed to slides with rubber cement. A rather thick layer of undiluted rubber cement was first spread on the slides. The Carbowax ribbons were added next. Lighter-fluid, essentially petroleum ether which can be substituted for it, was then run under the sections to dissolve the rubber cement and to float the ribbons. This notation medium did not dissolve the Carbowax and the ribbons could be manipulated in it for accurate location. The slides were dried on a 45° C warming table which also helped to flatten the sections. Adhesion was best when drying times were held to 4 hr or less. All excess rubber cement was washed away with xylene immediately prior to covering and the cover slips were carefully applied with a very thin resinous mounting medium to prevent dislodging the sections. Both aqueous and alcoholic stains have been used successfully and the slides have been left in them for as long as 3 days without loss of sections. The method was developed for fluorescence microscopy but serves equally well for visible light microscopy. Slides stained with a safranin-fast green combination have been used for both purposes, the safranin staining and fluorescing in a manner similar to rhodamine B.     

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.     

Stains recently certified

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.     

Preparing Serial Sections of Mature Corn and Wheat Kernels

Methods are described for preparing serial sections of paraffin-embedded mature corn and wheat kernels. Prior to embedding corn kernels are killed and fixed in formalin-aceto-alcohol (FAA), then steeped 5 days in 50% glycerol. After embedding by a special procedure, a thin slice is cut from one side of the kernel and the first few cell layers removed. The exposed surface is submerged in 20% glacial acetic acid in 60% ethanol for 2 or 8 days depending on the surface exposed, 2 days in air at 100% relative humidity at room temperature, and 2 days in air at 100% relative humidity at 8°C, successively. Wheat kernels, fixed in formalin-aceto-alcohol and embedded by the regular paraffin procedure, are similarly trimmed to expose a surface which is submerged in 20% glacial acetic acid in 60% ethanol for 2 days, 2 days in air at 100% relative humidity at room temperature and 2 days in air at 100% relative humidity at 8°C, successively. The corn and wheat kernels prepared by these methods give good serial sections when cut as thin as 14μ. The application of these methods to other seeds and caryopses is suggested.     

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).     

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.     

Carbowax 400: A New Solvent For Oil Red O And Sudan Iv For Staining Carbowax-Embedded And Frozen Sections

A method utilizing Carbowax 400 as the solvent for oil red O and Sudan IV is presented. This solvent fulfills all the requirements of an efficient solvent for lipid staining postulated by Chiffelle and Putt (1951) and, in addition, has several other advantages making it more practical than propylene glycol and ethylene glycol: (a) Carbowax sections are stained 3 times as fast, (b) frozen sections are stained 5 times as fast, (c) the staining solution is more readily prepared, and (d) the intensity of staining is greater.     

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.     

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.     

Diethylene Glycol Distearate Embedding and Ultramicrotome Sectioning for Light Microscopy

Diethylene glycol distearate can be used as an embedding medium for light microscopy. Two infiltration changes of about 6 hr each in the melted wax (melting point 47-52 C) are required before the final embedding which is done in 00 gelatin capsules for sectioning in the ultramicrotome by the procedure used in electron microscopy. Serial sections 1-2 μ thick can be cut without difficulty. No cooling devices are necessary for trimming and sectioning at laboratory temperature. Sections rarely become detached from the slides. The staining characteristics of the tissues are the same as when embedded in paraffin. For fluorescence microscopy, essentially the same procedure is followed. Tissues are not distorted and the intracellular structures are well preserved.