The Use of Papain in Clearing Plant Tissues for Whole Mounts

Materials killed and fixed in FAA (formalin-acetic acid-alcohol) and similar fixatives frequently are difficult to clear for whole mounts because the denatured proteins will not become soluble in NaOH and other clearing agents. If tissues are washed for 3 days in running water, then incubated at 40 C for 5-7 days in 2% papain buffered to pH 7.2 and activated with 15 ml of .02 M Na₂S, cell contents are partly digested. Normal clearing with 5-10% NaOH followed by chloral hydrate (sat. aq.) can then effect complete solublility of cell contents and their removal. Permanent slides can be made after staining (1% safranin O in 50% alcohol for 12 hr is successful), by dehydration through alcohols, clearing in xylene, and mounting in resin.     

Staining Nerve Fibers After Sublimate-Acetic and After Bouin'S Fluid

A silver staining method for paraffin sections of material fixed in HgCl₂, sat. aq., with 5% acetic acid is as follows. Process the sections through the usual sequence of reagents, and including I-KI in 70% alcohol, thiosulfate (5% aq.), washing and back to 70% alcohol containing 5% of NH₄OH (conc. aq.). After 3 minutes in the ammoniated alcohol, wash through tap water and 2 changes of distilled water and silver 5-10 minutes at 25°C. in 15% AgNO₃ aq. to which 0.02 ml. of pyridine per 100 ml. has been added. Blot the slide, but not the section and do not rinse. Reduce at 45°C. in 0.1% pyrogallol in 55% alcohol, then rinse in 55% alcohol and wash in water. The remainder of the process consists of gold toning, intensifying in oxalic acid, fixing in 5% Na₂S₂O₃, washing, dehydrating, clearing and covering. When the specimen contains much smooth muscle, the I-KI solution is acidified before use by adding 2 ml. of 1N nitric acid per 100 ml., and the sections treated for 3 minutes instead of the usual 2 minutes. Formalin should not be added to sublimate-acetic, but specimens that do not contain strongly argyrophilic nonneural tissue may be fixed in formalin or, preferably, Bouin's fluid. Sections of tissue after the latter type of fixation will not require the I-KI and thiosulfate but can go from 95% alcohol to the ammoniated alcohol. The advantages of fixing in HgCl₂-acetic acid are suppression of the staining of connective tissue and intensifying the staining of nerve fibers.     

Squash Method for Meiosis in Ovules

The ordinary Feulgen or acetic-lacmoid squash tech-nic following fixation in freshly made Carnoy's fluid (alcohol, 6: chloroform, 3: glacial acetic acid, 1), provides an easy and reliable method of studying meiosis in ovules. After fixation for 1 day, the material was hardened in 95% ethyl alcohol for 1-2 days and taken to water by gradual hydration. For staining by the Feulgen method, the material was hydrolyzed 8-10 minutes in 1 N HO at 58-60°C., followed by staining in decolorized leuco basic fuchsin for 2 hours. The staining was intensified by transferring the material to water. After 15-20 minutes the water was replaced by 45% acetic acid. For staining by acetic-lacmoid, the ovules after fixation, hardening and hydration were transferred to standard acetic-lacmoid stain to which was added 1 drop of 1 N HCl to every 10 drops of stain. Gentle heat was applied till the stain started to give fumes. After allowing 20 minutes at room temperature the material was transferred to fresh acetic-lacmoid. Some 6-12 ovules were mounted either in a drop of 45% acetic acid or acetic-lacmoid, depending upon the Feulgen or acetic-lacmoid staining respectively. Gentle and repeated tapping over the cover glass by a blunt needle loosened the cells of integument and nucellus and finally left the megaspore mother cells undergoing meiosis, fully exposed to view. The process was carried out under constant observation using the low power of the microscope. The desired amount of flattening was brought about by light pressure over the cover glass and gentle heating. The preparations were made permanent by dehydrating in ethyl alcohol and mounting in Euparal.     

A Clearing Technique for Detecting the Fungal Endophyte Acremonium sp. in Grasses

Leaf tissue of tall fescue Festuca arundinacea Schreb., hard fescue Festuca ovina L., red fescue Festuca rubra L. and perennial ryegrass Lolium perenne L. was stained with rose Bengal or aniline blue to detect the presence of the fungal endophyte Acremonium sp., Specimens were cleared using methyl salicylate, an optical clearing agent, and viewed using bright field microscopy. Tissue was presenred as dried tissue or stored in 70% aqueous ethyl alcohol before staining and clearing. Tissue was observed at 2, 4 and 12 weeks following clearing to check for stain retention. Staining with rose Bengal was inferior to aniline blue when followed by the clearing agent methyl salicylate. Fungal mycelia stained lighter with rose Bengal and were more difficult to detect than mycelia stained with aniline blue. The results illustrate the usefulness of combining staining and methyl salicylate clearing for detecting fungal endophytes.     

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.     

Amyl Acetate as a Clearing Agent for Embryonic Material

Amyl acetate is soluble in 95% alcohol and hot paraffin and produces no hardening in objects exposed to its action for prolonged periods. It may be advantageously employed as a general clearing agent and is especially recommended for refractory material. The following schedule has proven satisfactory for whole frog embryos and young tadpoles: 45 minutes to 1 hour in 95% alcohol, 24 hours in amyl acetate, rinse in toluene, 15 minutes each in three changes of paraffin, imbed. Material so treated may be sectioned at 5 μ with comparative ease.     

Water-Pretreatment Squash Technic: A New and Simple Practical Method for the Chromosome Study of Animals

To simplify the staining of animal chromosomes (especially in insect testes) the authors have borrowed (with necessary modifications) the squash technic of plant cytology. The method has four steps: (1) Water pretreatment. This step requires only about 5-10 minutes either in water at room temperature or in water kept at about 38°C. in a water bath. (2) Fixation. Ordinarily only 5 minutes in 10-15% aqueous solution of glacial acetic acid is necessary. (3) Staining. The fixed tissue is rinsed in two or three changes of distilled water and then placed in a solution of basic fuchsin: either 1% in 30% ethyl alcohol, or 0.2-0.4% in 5-10% lactic acid. In the former solution the staining period should be about 2 minutes: in the latter, 5-20 minutes. The time is not critical. (4) Squashing. The material is rinsed in several changes of distilled water, placed on a clean slide and squashed under a cover glass. Such preparations last 4-5 weeks, and a technic is described for removing the cover glass in order to mount in Euparal and to make them permanent. The authors list various species of vertebrates as well as invertebrates in which the technic has given good chromosome staining, as shown by illustrations.     

Alcoholic Thionin Counterstaining Following Golgi Dichromate-Silver Impregnation

Brains of cats that had been fixed 2 months or longer in 10% formalin were cut into 3-6 mm. slices and impregnated by Golgi's dichromate-silver procedure (6% dichromate solution, 4-6 days; 1.5% silver nitrate solution 2 days). Sections 100 µ thick were cut after embedding in low melting point paraffin. Three changes of xylene and three of absolute alcohol were followed by staining 3-5 minutes in a saturated solution of thionin in absolute alcohol. The sections were dipped quickly in absolute alcohol and cleared in xylene, then differentiation was effected by an equal-parts mixture of absolute alcohol and xylene. A final clearing in three changes of xylene and mounting in Permount completed the process. Counter-staining was most successful when applied to freshly cut sections.     

A Universal Stain for the Sex Chromatin Body

For the demonstration of the sex chromatin body in human tissues, fixation in 95% alcohol or modified Davidson's solution (95% alcohol, 30; formalin, 20; glacial acetic acid, 10; distilled water, 30) was best. The staining procedure chosen for most materials is the following: Mounted preparations are coated with celloidin, hydrated, hydrolyzed 20 min in 52V HCl at 20-25°C, rinsed thoroughly in several changes of distilled water and transferred to a buffered thionin solution. This consists of 3 parts: (1) A saturated solution of thionin in 50% alcohol (filtered); (2) Michaelis buffer: sodium acetate (3 H₂O), 9.714 gm; sodium barbiturate, 14.714 gm; CO₂-free distilled water, 500 ml; and (3) 0.1N HCl. To make the staining solution, mix 28.0 ml of the buffer solution with 32.0 ml of 0.1N HCl and bring the total volume to 100.0 ml with the thionin solution. Its pH should be 5.7 × 0.2, and care should be exercised that no acid is carried over from the hydrolyzing solution, since this would progressively lower the pH. The staining time varies from 15 to 60 min, depending on the specimen, but the shortest time consistent with adequate staining gives the clearest preparations. Slides are rinsed in distilled water and 50% alcohol and allowed to remain in 70% alcohol until the heavy clouds of stain cease to appear. Differentiation is completed in 80% and 95% alcohol, followed by dehydration in absolute alcohol, clearing in xylene and applying a cover glass with a synthetic resin (G. T. Gurr's DePeX was used). The sex chromatin is deep blue-violet and sharply contrasted against the lightly colored particulate chromatin of the nucleus. Cytoplasm remains unstained but fibrin and related structures show metachromasia. Chromosomes are well demonstrated if present. The method works on all types of tissues, is simpler and quicker than the Feulgen method, and often yields superior results.     

A Rapid Phloxine-Methylene Blue Oversight Stain for Formalin-Fixed Material

A rapid method is described, which yields brilliant and selective differential staining with phloxine-methylene blue. Aqueous phloxine (1%) is treated with hydrochloric acid (1 ml/l gm of phloxine) and the water-insoluble precipitate is washed, dried, and dissolved in acidified 95% ethyl alcohol. Slides are stained for several minutes in this solution (0.2%) followed by brief staining in the usual azure-methylene blue solution and differentiated with colophonium-alcohol. The method eliminates the necessity for prolonged staining at elevated temperatures, reducing the total staining procedure to approximately 15 min. In addition, controlled differentiation with colophonium-alcohol can be carried out on formalin-fixed material without loss of phloxine. The selectivity and tinctorial attributes of phloxine are also considerably improved.     

Staining Raw and Cooked Apple Tissues for Study of Cell Wall Structure

Permanent preparations were made of paraffin sections from raw and cooked apple tissues stained with microchemical color reagents for pectins and pentosans. Sections stained with ruthenium red to show pectins were dehydrated and covered in balsam, and sections stained with diphenylene diamine acetate (DDA) to show pentosans were washed with water and covered in Clearcol.

Cooking was accomplished by steaming cubed histological samples. Both raw and steamed specimens were fixed in FAA in a vacuum chamber, dehydrated and cleared in tertiary butyl alcohol, and embedded in paraffin. Paraffin sections first fixed to slides with Haupt's adhesive were further stabilized by immersing in a 1% celloidin solution after dissolving the paraffin.

Ruthenium oxychloride flakes were dissolved in a Coplin jar of water containing 2 drops of ammonium hydroxide. Rehydrated sections were stained in ruthenium red 30 minutes and rinsed in water. Three methods of further preparation follow: (1) Flood sections with 10% gum arabic; drain and air-dry thoroughly; immerse in xylene 5 minutes; cover in balsam. (2) Drain and air-dry sections; if desired, counterstain dry sections with Johansen's fast green solution; immerse in xylene; cover in balsam. (3) Dehydrate by dipping in 70%, 95%, and absolute ethyl alcohol; immerse in xylene; cover in balsam.

DDA was made by heating 15 g. of benzidine in 150 ml. of glacial acetic acid and 450 ml. of water until dissolved, then adding water to make 750 ml. of solution. Rehydrated sections were stained 4 hours in DDA, washed, stained 5 minutes in Congo red (Congo red, 5 g.; NaOH, 5 g.; water, 100 ml.), washed, and covered in Clearcol.

An Autotechnicon was used for dehydration, clearing, infiltration, deparaffinizing sections, and staining. Procedures that necessarily remained manual were fixation in a vacuum chamber, and all operations that followed staining.

Ruthenium red, though the best available indicator for pectins, may not be specific for these substances. DDA and ruthenium red stained identical structures in hypodermis and cortex. DDA also stained cuticle, hence was more useful than ruthenium red for delineating that portion. DDA sections were better for photomicrography, and for measuring thickness of cell walls. Neither stain prevented the study of cell walls in polarized light.     

A Method for Injecting Insect Tracheae Permanently

By the use of an alcohol insoluble dye (trypan blue), acetic acid, and a detergent (“Santomerse No. 3”), a resulting dye solution is obtained which will completely penetrate the tracheal system of an insect. The dye is injected by the use of a vacuum and by the pressure produced when the air is allowed to re-enter the dye vessel. The dye itself is permanently fixed in the tracheae by means of a fixing solution containing alcohol, acetic acid and barium chloride as its components. The material must be properly preserved after staining. It may be stored indefinitely in 70% alcohol, xylol, cedar oil or clove oil, depending on whether the material is to be used for sectioning or for whole mounts. Injected material may be sectioned in either celloidin or paraffin, or may be cleared and mounted in toto.     

Basic Two-Dye Stains for Epoxy-Embedded 0.3-1 μ Sections

Dyes used in the 3 methods recommended are: I, thionin and acridine orange (T-AO); II, Janus green and Darrow red (JG-DR); III, methyl green and methyl violet (MG-MV). The first 2 methods were two-solution stains, applied in sequence; the third, required only one solution since methyl violet is present in commercial methyl green. Staining solution and timing was as follows: Method I. 0.1% thionin in a 45% ethanolic solution of 0.01 N NaOH, 5 min at 70 C; rinsing in water and followed by 1 min in a 1% aqueous solution of acridine orange made up in 0.02 N NaOH, also at 70 C, then washed, and dried on slides. Method II. 0.5% Janus green in aqueous 0.05 N NaOH, 5 min at 70 C; rinsing in water then into 0.5% Darrow red in 0.05 N NaOH (aq.), 2 min at 70 C., washing, and drying on slides. Method III. 1% methyl green (commercial, unpurified) in 1% aqueous borax for 15-20 min at 20-25 C, washing and attaching to slides. All staining was performed by floating the sections on the staining solutions, all drying at 70 C, and mounting in a resinous medium. T-AO gave blue to violet cytoplasmic structures, darker nuclei which contrasted strongly with yellow connective tissue and the secretion of goblet cells. JG-DR resembled a hematoxylineosin stain, but by shortening the staining time in DR to 0.5-1 min, collagenous and elastic tissue retained more of the green dye. MG-MV gave dark green nuclei in light green cytoplasm, with collagenous and elastic tissues in blue to violet. As with most methods for staining ultrathin sections, thicknesses of less than 1 μ required longer staining times.     

Streptococcal Desoxyribonuclease for the Removal of Feulgen-Stainable Material

Removal of Feulgen-stainable material from the cell nucleus was accomplished by treatment of sections with streptococcal desoxyribonuclease. The procedure recommended is (1) Deparaffinize with xylene, followed by descending grades of alcohol. (2) Wash in tap water. (3) Treat slides for 1 hour at 37°C. with streptococcal desoxyribonuclease (1000 units/ml.) in 0.025M veronal buffer of pH 7.5 containing 0.003M MgSO₄. Treat control slides for an equal length of time at the same temperature. Renew the enzyme approximately every 15 minutes. (4) Wash slides briefly in tap or distilled water. (5) Dehydrate, then coat the sections by dipping in a 1% solution of celloidin in alcohol-ether. (6) Subject the preparations to the Feulgen reaction. Control slides showed characteristic nuclear staining; enzyme treated slides did not stain.     

Cytochemical Staining of Sections from Plastic-Embedded Flagellates

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

Histochemical Use Of Lead Tetra-Acetate I. Cleavage Of 1-2 Glycols

Lead tetra-acetate acts specifically to split the carbon-carbon single bond of the 1,2-glycol linkage to produce aldehyde radicals which may then be demonstrated by means of leucofuchsin, 2,4-dinitrophenlyhydrazine, or p-nitrophenylhydrazine. Routinely prepared slide sections from tissues fixed in 10% formalin are run down to 95% alcohol, rinsed in glacial acetic acid and then treated for 2 minutes in a saturated solution of lead tetra-acetate in glacial acetic acid with 5 g. of potassium acetate added for each 100 ml. of reagent. The sections are then washed in distilled water and placed in leucofuchsin for 10 minutes, or in a saturated 30% alcoholic solution of p-nitrophenylhydrazine for 5 minutes or 2,4-dini-trophenylhydrazine for 30 minutes. After staining, the sections are rinsed in 30% alcohol if the nitrophenylhydrazines were used, or in the standard dilute sulfite bath followed by running tap water for 5 minutes if leucofuchsin were used. Sections are routinely dehydrated, cleared, and covered. On examination, the sites of 1,2-glycol linkages will be stained violet by leucofuchsin or yellow by the nitrophenylhydrazines.     

Identification of Haemoglobin by its Catalase Reaction with Peroxide and o-Dianisidine

o-Dianisidine (3,3'-dimethoxybenzidine) when used as an indicator in the peroxidase activity of haemoglobin forms a clear and distinct orange stain. The reaction takes from 5 to 15 rain in the presence of H₂O₂ and completes both fixation and staining. This may be followed by dehydration in dioxane, clearing in xylene and mounting in D.P.X. or Canada balsam. The detergent Tepol can be used to spread or to disintegrate the chick embryo to obtain a monolayer of cells after the staining reaction has been completed. Background staining is negligible, the reaction is very sensitive and the colour developed is permanent Stock solutions are: (1) o-dianisidine, 100 mg per 70 ml of ethanol; (2) acetate buffer, 0.1 M at pH 4.6; and (3) hydrogen peroxide, 30% aqueous solution. The stock solutions should be refrigerated. The staining mixture contains o-diarusidine solution, acetate buffer, distilled water and hydrogen peroxide in the proportion of 4:1:1.5:0.2.     

Adaptation of Mallory's Trichrome Stain to Embryonic and Fetal Material

Based upon results of an investigation of the role of phosphotungstic acid in connective tissue staining, the Mallory trichrome stain was adapted to sequential application of all three dyes, thus making it usable on embryonic and fetal material. Ten to twelve day postconception mouse fetuses were formalin fixed and paraffin embedded. Staining was as follows: (1) 1% aqueous acid fuchsin for 5 min followed by not more than 30 sec in running tap water; (2) 2% aqueous phosphomolybdic acid (PMA) for 10 min followed by a 2 min running tap water wash; (3) staining in 0.5% aniline blue in 8% acetic acid for 10 min, followed consecutively by 30 sec in running tap water, 2% aqueous PMA for 2 min, and 30 sec in running tap water; (4) 2% orange G in 8% acetic acid for 5 min, and rinsing for 30 sec in running tap water. Dehydration in ethanol, t-butanol, acetone, or by blotting followed by 1:3 terpineol-xylene, clearing in xylene and mounting, completed the procedure. The 30 sec tap water rinses can optionally be replaced by 1-2 min in 8% acetic acid. Sections can be made redder by increasing acid fuchsin staining time, or increasing time in the first PMA; red can be decreased by decreasing staining time, increasing time of the 2 min tap water wash, or decreasing time in the first PMA. Blue or orange staining can be increased or decreased by varying staining times in these solutions. Sharper differentiation may be obtained by increasing the time in PMA.     

Staining Plant Tissues with Chlorazol Black E and Pianese III-B

The staining schedule was developed for a study of the mycorrhizae of red pine, Pinus resinosa Ait. From 70% alcohol, sections are stained in a saturated solution of chlorazol black E in 70% alcohol, 10-30 min; free dye removed by washing in 95% alcohol; stained 18-24 hr in Pianese III-b; rinsed in 95% alcohol, acidified by the addition of 2 ml of saturated aqueous picric acid per 100 ml, 3-4 changes or until the last change is pale yellow or light green; and rinsing in 95% alcohol to remove the acid. If the acid fuchsin is too intense, a cautious differentiation with 95% alcohol containing 1-3% of a 0.1 N solution of NaOH is made. If too much chlorazol black is removed, the effect can be compensated by overstaining with this dye at the beginning of the process. Sections are dehydrated, cleared, and covered in the usual manner. This stain has applications to plant tissues generally, and is particularly effective for meristematic tissues. It shows details of cytoplasmic structures and gives sharp delineation of primary cell walls.     

The use of basic Fuchsin in Plant Anatomy

A method useful in tracing the vascular bundles of herbaceous plants and the water conducting system of woody plants is presented. A stock solution of 2.5% basic fuchsin in 95% alcohol is prepared. A mixture of 1 to 2 parts of the stock solution to 100 parts of tap water (slightly alkaline) is made, standing 12-48 hours and being filtered before use. Cut ends of the living plants to be examined are immersed in the working solution until the dye appears in the parts desired.

The material may be examined by serial, hand sections mounted in 50% to 75% glycerin, or it may be split, laid open, and examined with a low power lens. Clearer views may be procured by dehydrating in an alcohol series and clearing in methyl salicylate. Permanent mounts may be made by removing the methyl salicylate with xylol and mounting in balsam. Structures such as leaves, which contain considerable chlorophyll, should be first boiled in slightly alkaline 85% alcohol until the green color is removed.