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.     

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     

A Buffered Giemsa-Bodian Stain for Neurological Material

A buffered Giemsa counterstain for the Bodian method is described. It is very useful for bringing out Nissl substance and nerve fibers in the same section. Bouin perfused or formalin fixed material from mammals, amphibia, reptiles and fish was used. After fixation, all tissues were decalcified for at least a week in 50% formic acid (1 part) and 20% sodium citrate (1 part). This was washed out thoroughly. The method described by Bodian (1936) was followed except for the following minor changes: Winthrop Protargol (Strong Protein Silver) Batch N346BJ was used exclusively; all glassware was cleaned with acid prior to setting up the stain; before developing, the sections were washed in warm tap water for 10 minutes; the gold chloride was chilled before use. The sections were then put into buffered Giemsa for 24 hours. Stock Giemsa: 0.75 grams powdered Giemsa (Coleman and Bell, Certification No. CGe-3) was dissolved in 50 ml. of glycerin overnight in the oven, then 50 ml. of methyl absolute alcohol were added. To 3 ml. of Giemsa stock solution, 87 ml. of distilled water buffered to pH 5.3 with 10 ml. of Sorensen's buffers were added and the solution filtered. Coleman buffer tablets gave best results at pH 5.0. Sections were then rinsed in 95% alcohol, two changes of absolute alcohol, two changes of xylene, and were then mounted in Clarite.     

Blood Serum as an Adhesive for Paraffin Sections

Diluted human and other mammalian blood serum (15 ml. of fresh blood serum diluted with 10 ml. of freshly distilled water, with 6 ml. of 5% formalin solution in distilled water added) can be used as a good adhesive for paraffin sections. It is preferable to Mayer's egg albumin-glycerol mixture because it is easily obtainable, can be quickly prepared, and sections are less subject to loosening after its use.     

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.     

Application of Methods for Mammalian Chromosome Spreads to the Cells of Cestodes

Tapeworm cells obtained by physical maceration between ground-glass surfaces are incubated for 3 hr in Hanks' balanced salt solution (BSS) supplemented with colchicine to a concentration of 10-⁴ M. After washing in BSS, the cells are incubated for 10 min in 1/4 strength BSS then centrifuged 10 min. Fixation of the intact button of cells (or alternatively, by squirting the cells directly into the fixative) in Carnoy's alcohol-chloroform-acetic acid (6;3:1) for 30 min follows, and cells, dispersed and washed in the fixative, are flattened by dropping the suspension on clean, water-wet slides which are then air-dried and stained with Giemsa diluted 1 ml;47 ml with distilled water to which 2 ml of buffer—M/15 KH₂PO₄, 32 ml, mixed with M/15 Na₂HPO₄, 68 ml—is added. After staining 15 min and washing in distilled water, slides are air-dried and mounted with resin. Well separated and well stained chromosomes have resulted.     

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

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

A 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.     

Staining Cells on a Membrane Filter with Giemsa Stock and Tap Water

Cells grown on type AM 6 Alpha Metricel Gelman filter membranes are fixed 1-2 min with OsO₄ vapor, washed in distilled water, dehydrated 4 min in 95% and 4 min in absolute ethanol. They are then stained for 2 min in a 1:1 dilution of Giemsa stock solution and an approximately neutral, low mineral content tap water. The stain is removed from the filter by 1 min in 50%, 1 min in 95%, 1 min in absolute ethanol and 1 min in a mixture of absolute ethanol and xylene 1:1. The filter is finally cleared in xylene and mounted with a synthetic resin. This procedure gave polychrome staining on neonatal thymic cells from C57 black pedigreed mice and on human leucocytes, with little or no stain retained by the filter membrane.     

Staining the Nucleus in Yeasts

Yeast cells kept young by repeated subculturing were centrifuged, washed twice in distilled water and smeared on slides coated with a little egg albumen. The cells were treated with 0.002 M 8-hydroxyquinoline for 1 hr, fixed first in OsO, vapour for 30 sec and then in chloroform for 30 sec. The slides were passed through descending grades of alcohol, washed in distilled water, then immersed in 0.17 M NaCl solution for 2 hr. at 57°C. They were again washed in distilled water and later hydrolysed in 1 N HCl at 60°C for 5-7 min. This was followed by washing in distilled water and in buffer. The slides were then kept for 3 hr in Giemsa stain comprising 96 ml of phosphate buffer of pH 7.0 and 4 ml of the stain. After dehydration, mounting was done in balsam. Nuclei were brightly stained and well differentiated; centrosomes were clear, and the process of nuclear division and movement to daughter cells could be studied. Pretreatment with 8-hydroxyquinoline increased the viscosity of the cytoplasm, while NaCl treatment and acid hydrolysis led to the complete removal of ribonucleic acid and basophilic material. A selective staining of chromatin was thus achieved. Structures resembling chromosomes could be seen when fixed and stained cells were squashed, soon after the removal of the slides from the stain, under a cover glass by applying uniform pressure with a rubber stopper. Fixation in osmic acid vapor and chloroform followed by acid hydrolysis and staining in leucobasic fuchsin also helps to obtain bright staining of the nucleus; however, the preparations are inferior to those obtained after 8-hydroxyquinoline, NaCl treatment and Giemsa staining.     

Pyronin Y in the Methyl-Green-Pyronin Histological Stain

Two samples of pyronin Y were found which, with the exception of eosinophilic granules and osteoid, stained only nucleic acids in animal tissues. Good differentiation was obtained. with n-butyl alcohol. It was therefore possible to prepare a differentially staining mixture of either of these pyronins combined with methyl green. This mixture stains polymerized desoxyribose nucleic acid (DNA) clear green, depolymerized DNA and ribonucleic acid red. The red staining of eosinophilic granules and osteoid is readily distinguished by its persistence after ribonuclease or warm-buffer extraction. The staining mixture consists of: (1) pyronin Y (Edward Gurr or G. T. Gurr), CHCl₃ extracted, 2% aq, 12.5 ml; (2) methyl green, CHCl₃ extracted, 2% aq, 7.5 ml; (3) distilled water, 30 ml. The staining procedure is as follows. (1) Immerse slides 6 min in the dye mixture. (2) Blot with filter paper. (3) Immerse in 2 changes of n-butyl alcohol, 5 min each. (4) Xylene, 5 min. (5) Cedar oil, 5 min. (6) Apply Permount and cover.     

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.     

An organic coating keeps orb‐weaving spiders (Araneae,Araneoidea, Araneidae) from sticking to their own capture threads

More than 95% of orb‐weaving spider species ensure prey capture success by producing viscous threads equipped with gluey droplets. However, this trap may bear serious risks for the web‐inhabiting spider as well. The obvious question, how a spider avoids getting stuck in its own capture spiral, has gained little scientific attention up till now. In 1905, the French naturalist Jean‐Henry Fabre concluded from anecdotal observation that orb‐weaving spiders protect themselves by a fatty surface coating. Here, we test this hypothesis by indirectly measuring the force necessary to detach an autotomized spider’s leg from the capture spiral of its own web (here called ‘index of adhesion’, IOA). Three groups of legs, each of the species Araneus diadematus Clerck, 1757 and Larinioides sclopetarius (Clerck, 1757), were tested. One was left untreated, one was washed with distilled water (H₂O), and one was washed with the organic solvent carbon disulphide (CS₂). In both species, we found a weak IOA between the spider leg and the gluey capture spiral in untreated and water‐washed legs without significant differences between the two. The IOA approximately doubled, when spider legs had been washed with carbon disulphide prior to measurement, that is, the CS₂‐washed legs stuck significantly more strongly than the untreated and water‐washed legs. These results provide indirect evidence for a protective anti‐adhesive organic coating on the spider’s body surface and so support Fabre’s hypothesis.     

Determination of Pseudomonas aeruginosa by Biochemical Test Methods

A new test method was devised for microbial gluconate oxidation, using an ammonium molybdate reagent. One loopful (about 2 mg wet wt.) of the test organism, grown on a nutrient agar plate for 18 hr, is transferred into 1 ml of the test liquid medium consisting of (NH₄)₂SO₄ 0.5 mg, potassium gluconate 10 mg, NaCl 5 mg, KH₂PO₄ 2 mg, MgSO₄·7H₂O 0.1 mg, and 1 ml of distilled water, incubated at 37 C for 6 hr without shaking, and then mixed with 3 ml of 1% aqueous solution of ammonium molybdate and 0.2 ml of glacial acetic acid. The mixture is heated in boiling water for 5 min, followed by abrupt cooling with running water. A deep blue colour appears in a positive result. A total of 39 strains of Pseudomonas aeruginosa showed positive results by this method, whereas Aeromonas, Vibrio, Proteus group, Klebsiella, Citrobacter and Enterobacter A group were all negative. Though some strains of Enterobacter B group showed a weak blue colour, it could be easily differentiated from the deep blue colour of Pseudomonas. Longer incubation of test microbes in the test medium, and longer heating of the reaction mixture gave unsatisfactory results.     

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.     

Laboratory Hints from the Literature: A Department Devoted To Abstracts Of Books and Papers From Other Joubnals Dealing With Stains and Microscopic Technic In General

Lesions produced in the cerebral cortex of rats were studied by Nauta's method for degeneration. The brains were perfused with physiological NaCl solution, followed by 10% neutral (CaCO₃) formalin. The brains were removed and stored in the formalin for 2 wk to 1 yr. Experimental modifications of the staining method showed that its sensitivity for fine degenerating fibers could he enhanced by the following changes: (a) omitting 0.05% potassium permanganate; (b) replacing the hydroquinone-oxalic acid mixture with 0.1% pyrogallol. Procedure: (1) frozen sections to water; (2) 0.5% phosphomolybdic acid, 45 min; (3) distilled water, 1 min; (4) 0.1% pyrogallol (aq.), 2 min; (5) distilled water, 3 washes of 1 min each; (6) 1.5% silver nitrate (aq.), 30 min; (7) distilled water, 1 min, (8) Laidlaw's ammoniated silver carbonate, 10110 sec; (9) Nauta's reducer, 1-2 min; (10) distilled water, 1 min; (11) 1.0% Na₂S₂O₃, 2 min; (12) distilled water 3 changes, 1 min each; (13) dehydrate, clear, and cover. This method gave equally good results on degenerating axons in both cortex and thalamus.     

Silver Protein Staining of Nerve Fibers in Celloidin Sections

Celloidin sections from formalin-fixed brain and spinal cord of primates are stored in 70% alcohol after cutting, soaked in 2% pyridine in 50% alcohol for 6-8 hr at 37 C, and transferred to 1% concentrated NH₄OH in 50% alcohol 15-18 hr at 20-25 C. After washing and flattening, the sections are transferred to 1% silver protein solution containing 30 ml of 0.2 M H₃BO₃/100 ml. Impregnation is accomplished in 50 ml screw-top jars, 50 mm in diameter, which are filled to a depth of 35 mm, and have 1 gm of copper foil, 0.002 inch thick added. The foil is folded in loose accordion-fashion, pierced and threaded, cleaned in 5% HNO₃, rinsed in distilled water, and suspended in the solution just above the sections by fastening the thread to the jar lid. The sections are impregnated for 24 hr at 37 C, rinsed in distilled water, reduced in a solution of 5% Na₂SO₃ and 1% hydroquinone for 10 min, washed in distilled water and toned in 0.2% gold chloride for 5 min. After rinsing in distilled water, the sections are transferred to 1% oxalic acid for 45-60 sec, washed in distilled water and placed in 5% Na₂S₂O₃ for 5 min. Sections are then washed, dehydrated to 95% alcohol, cleared in terpineol, followed by 3 changes in xylene, and mounted.     

The Peracetic-Orcein-Halmi Stain: A Stain for Connective Tissues

A selective stain useful for the study of connective tissues is described. The stain demonstrates elastic and oxytalan fibers as well as fibrils in mucous connective tissues previously undescribed. Reticular fibers are not stained. The stain may be used on sections that have been fresh frozen or fixed in formalin or ethanol. Sections are deparaffinized, washed in absolute ethanol, oxidized in peracetic acid 30 min, washed in running water, stained in Taenzer-Unna orcein 15 min, 37°C, differentiated in 70% ethanol, washed in running water, stained in Lillie-Mayer alum hematoxylin 4 min, blued in running water, and counterstained 20 sec in a modified Halmi mixture of 100 ml distilled water, 0.2 gm light green SF, 1.0 gm orange G, 0.5 gm phosphotungstic acid and 1.0 ml glacial acetic acid. Sections are rinsed briefly in 0.2% acetic acid in 95% ethanol, dehydrated and mounted.     

Permanent Mounting of Unstained and Aceto-Orcein Stained Cells in the Water-Soluble Medium,Abopon

For cellular morphology, mammalian cells were grown on cover slips in Leigh ton tubes, fixed in 1% osmic acid vapor for 2 min, decolorized with 30% H₂O₂ in 5% ammonium oxalate solution (1:7) for 2 min, then washed thoroughly, and finally mounted in a water-soluble medium consisting of a saturated solution of Abopon in 0.2 M phosphate buffer, pH 7.0. For chromosomal analysis of similarly cultured cells, aceto-orcein preparations were made by conventional methods, with the following minor modifications: following pretreatment with colchicine and hypotonic expansion, the cells on the cover slips were fixed in acetic-alcohol (1:3), air dried, incubated at 37° C for 15 min in 2% orcein in 45% acetic acid, rinsed in 45% acetic acid, washed several times in distilled water, and finally mounted in Abopon mounting medium. Both kinds of preparations were allowed to harden for 24 hr before being handled. Such slides will keep for years at room temperature. Studies requiring frequent comparisons of cellular and chromosomal morphology of cultured cells can thus be extended over long periods of time.     

Staining and Mounting Helminths

The following procedure is recommended: Fix ces-todes and trematodes (while held flat between glass slides) 0.5–2.0 hr. in the following mixture: formalin, 15; acetic acid (gl.), 5; glycerol, 10; 95% ethyl alcohol, 24; distilled H₂O, 46; all proportions by volume. After freeing them from the slides, wash thoroughly in running water and stain immediately thereafter. Stock staining solution: ferric ammonium alum (violet cryst.), 2 g.; distilled H₂O (cold) 100 ml.; after solution, add 2 ml. concentrated H₂SO₄, bring to a boil; add 1 g. coelestin blue B (Nat. Aniline), boil 3–5 min.; cool and add 10 ml. absolute methyl alcohol and 10 ml. glycerol. Dilute 1 vol. with 3 vol. distilled H20 for use. Stain 5–30 min., depending on size of specimens. Wash with 2 changes 0.5 hr. each of distilled H₂O, then 50% isopropyl alcohol 12–16 hr., 50% isopropyl alcohol 2 hr., followed by graded isopropyl alcohol for dehydration. Ether: ethyl alcohol (equal parts), 1 hr., is followed by embedding in celloidin in a sheet just thick enough to cover the specimens. Trim embedded specimens and dehydrate with isopropyl alcohol, 80%, 90% and absolute. Clear in beechwood creosote. Mount in balsam with cover glasses that overlap the edges of the celloidin 1–2 mm. While drying at 37°C, refill edges of mount with fresh balsam as needed. When dry, remove excess balsam and ring the edges with ordinary gloss enamel paint.