Adaptation of the Millon, Sudan Black B, and Periodic Acid-Schiff Technics for Block Staining of Insect Tissues

Spermatophores and reproductive systems of the beetle, Lytta nuttalli Say, fixed in Bouin's aqueous picroformol or buffered 10% neutral formol were stained in toto by the Millon, Sudan black B and periodic acid-Schiff reactions as follows. Millon: after excess fixative is removed in 70% ethanol, specimens are brought to water, stained in Millon's reagent at 60 C for 1 hr, rinsed in 2% aqueous nitric acid at 40-50 C, dehydrated rapidly, cleared, embedded and sectioned as usual. Sudan black B: specimens are taken to absolute ethanol, stained in a saturated solution of Sudan black B in absolute ethanol at room temperature for 24-48 hr, rinsed and cleared in xylene, embedded and sectioned. PAS: specimens are brought to water, oxidized in 0.5 aqueous HIO₄ at 37 C for 30 min, washed in 2 changes of water, stained in Schiif reagent at room temperature for 1 hr, rinsed in 3 changes of 0.5% aqueous potassium metabisulfite, washed in running water for 10-15 min, dehydrated, cleared, embedded and sectioned. All 3 methods produced their characteristic staining in specimens up to 3 mm thick     

Preparation of Cuticle-free Epidermal Peels for the Demonstration of Primary Pit-fields

Cuticle-free epidermal peels for the demonstration of primarypit-fields (if any) on the outer surface of the epidermal cells,can be prepared by a chemically controlled process, using 5-10per cent NaOH solution as a macerating medium. Such peels canbe obtained from any desired part of the plant within a periodof a week. The process involves allowing the trimmed piecesof the plant material to stand in the NaOH solution at 50–60°C, until all the cuticular depositions on the outer surfaceof the epidermal cells are dissolved and the unwanted tissuesare sufficiently loosened to allow the easy removal of the epidermis.The separation of epidermis is done by giving moderate agitationsin water during washing or by means of a soft-haired brush undera dissecting microscope. The peels so obtained must be washed thoroughly in water andcan then be stained with bismark brown or iron alum hematoxylln.After dehydration in ethanol, they can be mounted permanentlyin Canada balsam.     

CUTICLES OF TWO SPECIES OF ALETHOPTERIS

Specimens of Alethopteris sullivanti (Lesquereux) Schimper and Alethopteris lesqueieuxi Wagner were found in Middle Pennsylvanian coal balls from the Lovilia, Iowa and West Mineral, Kansas localities. Light microscopy and scanning electron microscopy (SEM) were used to study macerated cuticles of these two foliage types of the pteridosperm, Medullosa. The lower epidermises of both species of Alethopteris possess sunken stomata, papillae, and evidence of multicellular hairs. The upper epidermises are devoid of these features. Critical point drying and subsequent observation with SEM were used to document the structure of the multicellular hairs of A. lesquereuxi. Replicas of the lower epidermal features of an impression-compression specimen of A. sullivanti were observed with SEM and compared with the petrified specimens.     

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.     

Enzyme clearing of alcian blue stained whole small vertebrates for demonstration of cartilage.

Preparation of small vertebrates cleared after alcian blue staining of cartilage is facilitated by trypsin digestion. Specimens are fixed in formation, washed, skinned, and eviscerated. After staining in a solution of alcian blue in acetic acid-alcohol for 24-48 hours, they are transferred to water through graded alcohols. Excess alcian blue is removed over a period of up to three weeks by changes every 2-3 days of 1% trypsin in approximately one-third-saturated sodium borate. Bony tissues may be stained after this in a solution of alizarin red S in 0.5% KOH. Specimens are bleached if necessary and dehydrated through graded KOH-glycerine mixtures for storage in glycerine. Since alcohol treatment in addition to formalin fixation does not affect results with this method, it should be useful to researchers who want to study the cartilage or cartilaginous skeletons in museum specimens, which are routinely fixed in formalin and stored in alcohol.     

Enzyme Clearing of Alcian Blue Stained Whole Small Vertebrates for Demonstration of Cartilage

Preparation of small vertebrates cleared after alcian blue Staining of cartilage is facilitated by trypsin digestion. Specimens are fixed in formalin, washed, skinned, and eviscerated. After staining in a solution of alcian blue in acetic acid-alcohol for 24-48 hours, they are transferred to water through graded alcohols. Excess db blue is removed over a period of up to three weeks by changes every 2-3 days of 1% trypsin in approximately we-third-saturated sodium borate. Bony tissues may be stained after this in a solution of alixarin red S in 0.5% KOH. Specimens rue bleached if necessary and dehydrated through graded KOH-glycerine mixtures for storage in glycerine. Since alcohol treatment in addition to formalin fixation does not affect results with this method, it should be useful to researchers who want to study the cartilage or cartilaginous skeletons in museum specimens, which me routinely fixed in formalin and stored in alcohol.     

Staining of Animal Tissues with the Dye Base of Methylene Green in Benzene to Facilitate Identification and Selection of Material

Methylene green as the free dye base dissolved in benzene can be used to stain animal tissues sufficiently during clearing for paraffin embedding to distinguish morphological features. It can therefore be used to allow trimming off unwanted tissues, to orient specimens when casting in wax and to select sections from a wax ribbon. The stain is removed very quickly on contact with water, and has no effect on subsequent staining techniques. The dye base is prepared by adding 2.5 ml of 10% NaOH to 100 ml of 1% aqueous methylene green, mixing well and allowing to stand at 20-30 C for 24 hr. The precipitate is separated by filtration, washed on the paper and dried at 50-55 C. The dry precipitate is dissolved in 200 ml of benzene to make the stock solution, which is diluted with 1 or 2 parts of benzene for tissue blocks, but used undiluted on sections.     

A Modified Pyridine-Silver Stain for Teased Preparations of Motor and Sensory Nerve Endings in Skeletal Muscle

This technique has been developed especially to stain sensory receptors which have been localised intramuscularly by electrophysiological means. Rat intertransverse caudal muscles, removed immediately after death, are fixed for 24 hr in a freshly prepared mixture of absolute ethyl alcohol, 4.5 ml; distilled water, 5 ml; and concentrated HNO_a, 0.1 ml. After a further 24 hr in 10 ml of absolute ethyl alcohol containing 0.1 ml of ammonia solution (sp. gr. 0.88), the muscles are washed in distilled water for 30 min and placed in full strength pyridine for 2 days. They are then washed for 24 hr in distilled water (changed 5-8 times) and left in 2% AgNO₃, in the dark for 3 days at 25 C. Following reduction in 10 ml of 5% formic acid containing 0.4 gm of pyrogallol for 6-24 hr, the specimens are washed briefly in distilled water and stored in pure glycerol. The nerve endings can then be teased out and mounted in glycerol, under cover glasses ringed with a waterproof cement. The advantage of this method is that it gives consistently good staining of receptors and motor end-plates in small muscles of the rat     

Staining Mitochondria in Fixed Blood Smears

Wet blood smears are placed immediately in Helly's fluid for 24 hr, transferred directly to a saturated solution of potassium dichromate for 48 hr and washed in running water for 2-4 hr. The slides are then treated with iodine and sodium thiosulf ate and washed several hours or overnight. Excess water is removed by blotting the slide around the smear, Altmann's aniline fuchsin is placed on the smear and the slide is heated over a spirit lamp until white fumes appear. After the slide cools the stain is poured off and the excess removed by washing with distilled water. Methyl green (1% aqueous) is dropped on the smear and left for approximately 30 sec. It is then passed rapidly through 2 changes of absolute ethanol and into xylene, from which it is mounted in Permount. This stains mitochondria, red blood corpuscles and specific granules of eosinophilic granulocytes red on a green background.     

Erratum

A solution of plastic consisting of toluene, 720 ml; methanol, 180 ml; ethyl cellulose (Ethocel, standard 7 CPS), 250 gm; and Dow resin 276 V-2, 75 gm is applied to a leaf surface which has been dampened with toluene. The plastic is spread to a thin film with the edge of a card and allowed to dry. After drying, the plastic may be peeled from the leaf surface and either stored dry in a small envelope or mounted permanently on a microscope slide. Permanent mounts are prepared by placing a small section of the peel from the upper and lower surfaces of the leaf on a microscope slide and covering with a No. 1 cover glass. A small spot of balsam on each corner of the cover glass secures the glass in position. This air mount has proved to be superior to water, glycerol or balsam mounts. Fresh leaves are washed with a mild detergent before application of the plastic. Herbarium specimens are soaked in water overnight to restore the leaf to a semiturgid condition. Five species of different plant families have been illustrated to show the diagnostic features of the surface of the cuticle. An isolated layer of epidermal cells obtained by chemical maceration permitted cell and imprint comparison. The remarkable amount of detail shown by the prints is an aid for phylogenetic studies and may make the recognition of fossil cuticles possible.     

Surface Printing of Plant Leaves for Phylogenetic Studies

A solution of plastic consisting of toluene, 720 ml; methanol, 180 ml; ethyl cellulose (Ethocel, standard 7 CPS), 250 gm; and Dow resin 276 V-2, 75 gm is applied to a leaf surface which has been dampened with toluene. The plastic is spread to a thin film with the edge of a card and allowed to dry. After drying, the plastic may be peeled from the leaf surface and either stored dry in a small envelope or mounted permanently on a microscope slide. Permanent mounts are prepared by placing a small section of the peel from the upper and lower surfaces of the leaf on a microscope slide and covering with a No. 1 cover glass. A small spot of balsam on each corner of the cover glass secures the glass in position. This air mount has proved to be superior to water, glycerol or balsam mounts. Fresh leaves are washed with a mild detergent before application of the plastic. Herbarium specimens are soaked in water overnight to restore the leaf to a semiturgid condition. Five species of different plant families have been illustrated to show the diagnostic features of the surface of the cuticle. An isolated layer of epidermal cells obtained by chemical maceration permitted cell and imprint comparison. The remarkable amount of detail shown by the prints is an aid for phylogenetic studies and may make the recognition of fossil cuticles possible.     

Sonication and Other Improvements on Jeffrey's Technique for Macerating Wood

Large, up to pencil-size pieces of wood, rather than slivered wood, are macerated in Jeffrey's fluid, which consists of ca. 1:1 aqueous nitric acid and aqueous chromic acid (each 10% or weaker). Staining is done in ca. 1% solution of safranin in absolute alcohol. The intact pieces of wood can be very briefly sonicated in xylene to release isolated elements and connected cell groups. Various alternative steps are noted. Discussion focuses on the practical and theoretical advantages derived from the aforementioned innovations.     

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

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

The Histochemical Detection of Thallium

Thallium can be histochemically localized in formalin-fixed, paraffin-processed tissues by treating sections, after passing them to water through xylene and graded alcohols, in a 0.5% aqueous Bi(NO₃)₃ solution acidified with 1 drop of concentrated HNO₃ per 100 ml. Sections are then washed and placed for 5 min in 2% aqueous KI, again washed, routinely dehydrated, cleared and covered. If desired, they may be lightly counterstained prior to dehydration. The tissue sites of T1 will be demonstrated by the fine wine-red crystals of (T1I)₂-BiI₃ resulting from the reaction, and will appear black with the usual microscopic illumination.     

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 Phloxine-Azure-Hematoxylin Sequence for Differential Staining of Cells in Pancreatic Islets

Sections of 6 μ from tissues fixed in Susa or in Bouin's fluid (without acetic acid) and embedded in paraffin were attached to slides with Mayer's albumen, dried at 37 C for 12 hr, deparaffinized and hydrated. The sections fixed in Susa were transferred to a I₂-K₁ solution (1:2:300 ml of water); rinsed in water, decolorized in 5% Na₂S₂O₃; washed in running water, and rinsed in distilled water. Those fixed in Bouin's were transferred to 80% alcohol until decolorized, then rinsed in distilled water. All sections were stained in 1% aqueous phloxine, 10 min; rinsed in distilled water and transferred to 3% aqueous phosphotungstic acid, 1 min; rinsed in distilled water; stained 0.5 min in 0.05 azure II (Merck), washed in water; and finally, nuclear staining in Weigert's hematoxylin for 1 min was followed by a rinse in distilled water, rapid dehydration through alcohols, clearing in xylene and covering in balsam or a synthetic resin. In the completed stain, islet cells appear as follows: A cells, purple; B cells, weakly violet-blue; D cells, light blue with evident granules; exocrine cells, grayish blue with red granules.     

Histoautoradiographic Localization of Calcium in Oat Plant Tissues

The leaching of water-soluble and exchangeable calcium in histoautoradiog-raphy of oat tissue can be prevented by using acetone as the dehydration fluid (freeze substitution technique) and by keeping the tissue sections, while stretching on water, embedded in the methacrylate matrix. Ca⁴⁵ was either added to the mineral solution on which the oat plants were grown (75 μc), or applied on the leaf surface (8 μc). After freezing in melting isopentane, specimens of 1-2 mm dimensions are fixed for 24 hr in an acetone-OsO₄ (1%) solution at—80 C. Dehydration is obtained by transferring the material every day for 6 successive days to a fresh acetone solution at—80 C. The material is infiltrated by a three-time renewed monomer methacrylate mixture (methylmethacrylate I, butylmethacrylate 4) at—50 C. The specimens are embedded in the polymerizing methacrylate mixture at room temperature. Sections of 4-8 μ are easily cut with a rotating microtome. If the methacrylate is not removed from the sections, they can be stretched on water without leaching of calcium. The presence of methacrylate in no way hinders microscopic observation nor effective histoautoradiography.     

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

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

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

Evidence for the Uptake of Large Anions through Stomatal Pores

Experiments were conducted with leek (Allium porrum L.) leaves to investigate whether aqueous solutions are able to penetrate stomata. Epidermal strips were used for the determination of transport rates. Stomata were opened by fusicoccin or closed by darkness or abscisic acid. A droplet containing the anionic fluorescent dye, uranine, was placed on the physiologically outer side of the epidermis and allowed to dry. With open stomata 30 times more uranine penetrated through the epidermal strips than with closed stomata (comparison of medians). In another experiment droplets of uranine solution were placed on leaf segments and epidermal strips were removed after drying of the droplets. Penetration of uranine through stomata was detectable under the microscope both with epidermal strips from the transport experiments and with strips obtained after application on leaf segments. As maximum uptake rates occurred during the drying process, it is concluded that penetration took place via water films. These results show that the physical restrictions preventing stomatal penetration of static droplets are not decisive for drying droplets and that stomatal uptake of dissolved ionic substances occurs under natural conditions, i.e. without surfactants or applied pressure.     

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.