Possible uses of Dioxan in Botanical Microtechnic

Dioxan has been well established as an advantageous dehydrating agent for plant tissues. It dehydrates equally well after fixatives containing formalin, acetic acid, chromic acid, chromates, mercuric chloride, osmic acid, and alcohol. Better infiltration of paraffin after dehydration may be obtained by passing the material thru (1) a cold bath composed of 30 cc. of dioxan, 5 cc. of xylol and 20 cc. of melted soft paraffin and, (2) a warm bath of 50 cc. of dioxan, 50 cc. of paraffin, and 10 cc. of xylol. Transfer from (2) to soft paraffin. A dioxan fixative consisting of dioxan 50 cc., formalin 6 cc., acetic acid 5 cc., water 50 cc. was devised for delicate subjects. The fixed material is transferred directly into dioxan and mounted in dioxan-diaphane or dioxan-balsam. Very delicate objects require dioxan dilution of the balsam and slow concentration of the mounting medium by evaporation.

Entire plant parts or epidermal peelings are fixed in any desired fixative, washed if necessary, transferred to dioxan and mounted in diluted dioxan-balsam or diaphane. Dioxan may be used to mount hyalin objects whose refractive indexes approach those of balsam in media of higher index than balsam. It may be used in place of alcohol in finishing parafin sections, and since it exhibits different stain solubilities than alcohol it offers an important new tool in obtaining and maintaining stain balances.     

Some Staining Methods for Bacteria and Yeasts

For staining flagella of bacteria use actively motile organisms 20 to 24 hours old, allow to diffuse in sterile water 20 to 30 minutes, transfer droplets of the suspension to clean slides and let evaporate without spreading. Then treat 2 to 4 minutes with the following mordant: tannic acid 10 or 20%, 50 cc.; ferric chloride 5%, 10 to 15 cc.; carbol fuchsin (Ziehl-Nielson), 5 cc.; hydrogen peroxide 3%, 6 to 8 cc. Wash and stain 2 to 3 minutes with a mixture of basic fuchsin, saturated alcoholic, 10 cc.; anilin oil (1 part) and 95% alcohol (3 parts) mixed, 5 cc.; distilled water, 30 cc.; acetic acid, 4%, 1 cc. Wash thoroly with water.     

Staining Paraffin Sections with Protargol: 1. Experiments with Bodian's Method. 2. Use of Jvpropyl and N-Butyl Alcohol in Hofker's Fixative:

Paraffin sections of nervous tissue, which had been fixed in Hofker's fluid, stained readily with protargol solution without the addition of metallic copper or other activator. Amidolsulfite mixtures reduced the protargol more rapidly and completely than hydroquinone-sulfite. Intensification of the stain could be secured by reducing with 0.5% amidol (or pyrogallol) solution after gold toning. The completeness of staining of unmyelinated fibers of the dorsal roots of cat spinal nerves was checked by estimating the number of fibers in a root and the cells of its associated ganglion. A fiber cell ratio of 1:1 was found hi 4 specimens, indicating within limits of error that all fibers were stained. An improvement of die original Hofker's mixture as a fixative was obtained by using a mixture of formic acid, 5 cc.; trichloracetic acid, 10 g.; n-propyl alcohol, 20 cc.; and n-butyl alcohol, 60 cc. (instead of the acetic, trichloracetic, ethyl alcohol mixture used hi the original formula). The following arbitrary method is suggested. Fix 12 to 24 hours, pass to water thru graded ethyl alcohol, wash several hours, dehydrate and embed in paraffin. Cut, mount, and remove the paraffin, pass to water and impregnate 2 or 3 days at 27 to 30$$C. in a 0.5% aqueous solution of protargol (Winthrop Chemical Co.). Rinse 2 or 3 seconds and reduce with 0.5% amidol (Agfa brand used) in 5% sodium sulfite solution. Wash, tone with 0.1% gold chloride, wash and reduce with 0.5% amidol (no sulfite), wash, dehydrate and cover. The method works well on spinal nerve roots, cerebrum, cerebellum, and spinal cord, and moderately well on nerve trunks including sympathetic nerves. Tissues from cat and guinea pig were used.     

A Rapid Combined Fixing and Staining Method for Plant Chromosome Counts

A new method for rapidly preparing slides suitable for chromosome counts by the use of a combined fixing and staining solution involves the substitution of anthraquinone for picric acid in Bouin's formula and the addition of alizarin red S with a metallic salt as a mordant. The fixed smears, after being dehydrated to 95% alcohol, are differentiated in 0.5% sulfuric acid in 95% alcohol saturated with picric acid, washed, cleared and mounted in xylol-balsam. Cymene may be used to intensify the stain. Root tips fixed in the above solution may be dehydrated in dioxan, a paraffin solvent; infiltrated, embedded, sectioned and mounted in the usual way. The sections are subsequently differentiated in picro-sulfuric acid alcohol and cymene. An alternative method of differentiation for this stain is also described.     

A Comparative Study of some Dehydration and Clearing Agents

An experiment to determine the advantages of diozan, iso-butyl alcohol, tertiary butyl alcohol, and ethyl alcohol as dehydrants and chloroform, toluol, xylene, benzol, methyl benzoate, methyl salicylate, and acetone as clearers is described. Materials fixed in Bouin's fluid, Zenker formol, and 10% neutral formalin were dehydrated, embedded, sectioned, and stained. Bouin's fluid produces less hardening, shrinkage and distortion than the other fixatives employed. Slow dioxan is the best method of dehydration. All the picric acid need not be removed from tissues to be embedded in paraffin. Tissue blocks not more than 4 mm. thick may be dehydrated and impregnated with paraffin by slow dioxan in 13 hours, fast dioxan in 10 hours, iso-butyl alcohol and tertiary butyl alcohol in 14 hours, and ethyl alcohol-chloroform in 17 hours without incurring any distortion due to rapidity of dehydration and infiltration.     

Staining Sections of Peripheral Nerves for Axis Cylinders and for Myelin Sheaths

Pieces of mammalian nerves 1 to 2 cm. long were placed under moderate tension and fixed 24–48 hours in: picric acid, saturated aqueous, 90 ml.; formalin, 10 ml.; and trichloracetic acid, 25% aqueous, 2 ml. They were washed in water, cut in two and one end stained with 0.04–0.06% osmic acid solution, while the other was dehydrated, embedded in paraffin, and mounted sections from it stained with protargol. The fixing solution used was selected from a number of combinations of acidified picro-formalin as the one most likely to give satisfactory results when followed by both silver and osmic acid. The use of osmic acid solutions of less than 0.1% concentration avoided the overstaining of myelin sheaths seen frequently when stronger solutions were used with material that had been fixed previously. Protargol, 0.5% solution with fast green FCF added to make 0.05% dye in the final concentration, was used to impregnate sections for axis cylinders. Reduction and toning were done as in Bodian's method.     

Staining Sections of Peripheral Nerves for Axis Cylinders and for Myelin Sheaths

Pieces of mammalian nerves 1 to 2 cm. long were placed under moderate tension and fixed 24-48 hours in: picric acid, saturated aqueous, 90 ml.; formalin, 10 ml.; and trichloracetic acid, 25% aqueous, 2 ml. They were washed in water, cut in two and one end stained with 0.04-0.06% osmic acid solution, while the other was dehydrated, embedded in paraffin, and mounted sections from it stained with protargol. The fixing solution used was selected from a number of combinations of acidified picro-formalin as the one most likely to give satisfactory results when followed by both silver and osmic acid. The use of osmic acid solutions of less than 0.1% concentration avoided the overstaining of myelin sheaths seen frequently when stronger solutions were used with material that had been fixed previously. Protargol, 0.5% solution with fast green FCF added to make 0.05% dye in the final concentration, was used to impregnate sections for axis cylinders. Reduction and toning were done as in Bodian's method.     

A Urea Silver Nitrate Method for Nerve Fibers and Nerve Endings

A silver nitrate stain for nerve fibers and endings applicable to paraffin sections on the slide utilizes the properties of urea to accelerate the procedure and improve the specificity of the stain. After removal of the paraffin the sections are run through absolute, 95% and 80% alcohol and placed for 60-90 minutes at 50-60°C. in: 1% aqueous silver nitrate, 100 ml.; urea, 20-30 g.; 1g. mercuric cyanide and 1 g. picric acid in 100 ml. of distilled water, 1-3 drops. After the silver bath they are rinsed quickly in 2 changes of distilled water and reduced for 3-5 minutes at 25-30°C. in: water, 100 ml.; sodium sulfite, anhydrous, 10g.; hydroquinone, 1-2g.; urea, 20-30g. They are then washed thoroughly in 4-5 changes of distilled water, passed through graded alcohols into 80% alcohol and examined under the microscope. If nerve fibers are not distinct, the sections are returned to the same urea-silver-nitrate bath for 10-15 minutes, rinsed, reduced, washed and dehydrated as before. This process may be repeated until staining is adequate; then they are dehydrated, cleared, and mounted.

Nerve fibers show a color range from brown to black; nerve cells from yellow to brown; and the background, depending on the type of tissue and its fixation, from yellow to light brown.     

The Use of Salicylic Acid as a Fixative

The following salicylic acid-containing fixatives are useful for cytological studies in plants. The first, here designated HFC, is recommended for studies on somatic mitosis and chromosome individuality. The second, denoted HFP, is recommended for studies on plastids.

HFC is made up in two solutions. Sol. A: 100 cc. sat. aq. sol. salicylic acid, slight excess copper hydroxide, 20 cc. formaldehyde, 30 cc. normal ortho-phosphoric acid, 200 cc. water, 1 g. saponin; pH 1.8 to 1.9. Fix in Sol. A 15 to 30 minutes in partial vacuum of 35 cms. Then add Sol. B: 1% aq. chromic acid in equal parts. Continue fixation for period of 18 to 24 hours.

HFP is also made up in two solutions which are used in equal parts. Sol. A: 100 cc. sat. aq. sol. salicylic acid, slight excess copper hydroxide, 10 cc. normal ortho-phosphoric acid, 1/2 g. saponin. Sol. B: 187.5 cc. 1% aq. chromic acid, 50 cc. 2% osmic acid. Fixation technic as HFC.

Dehydrate and infiltrate with paraffin after Zirkle. Stain with crystal-violet-iodine.     

The Use of Salicylic Acid as a Fixative

The following salicylic acid-containing fixatives are useful for cytological studies in plants. The first, here designated HFC, is recommended for studies on somatic mitosis and chromosome individuality. The second, denoted HFP, is recommended for studies on plastids.

HFC is made up in two solutions. Sol. A: 100 cc. sat. aq. sol. salicylic acid, slight excess copper hydroxide, 20 cc. formaldehyde, 30 cc. normal ortho-phosphoric acid, 200 cc. water, 1 g. saponin; pH 1.8 to 1.9. Fix in Sol. A 15 to 30 minutes in partial vacuum of 35 cms. Then add Sol. B: 1% aq. chromic acid in equal parts. Continue fixation for period of 18 to 24 hours.

HFP is also made up in two solutions which are used in equal parts. Sol. A: 100 cc. sat. aq. sol. salicylic acid, slight excess copper hydroxide, 10 cc. normal ortho-phosphoric acid, 1/2 g. saponin. Sol. B: 187.5 cc. 1% aq. chromic acid, 50 cc. 2% osmic acid. Fixation technic as HFC.

Dehydrate and infiltrate with paraffin after Zirkle. Stain with crystal-violet-iodine.     

Simple Aids to Microscope Illumination

The following rapid but reliable method of making permanent preparations from temporary mounts has proved to be very useful.

Pollen mother-cell smears: Smeared anthers are treated hi the usual way with Belting's acetocarmine, except that the cover slip is left off. When correct differentiation is attained the stain is thoroly washed off with 50% acetic acid and the slide flooded with dioxan. This is followed by 2 changes of dioxan for 2 minutes each. A drop of Canada balsam dissolved in dioxan is added and a cover slip applied. In cases where a cover slip has been used at the acetocarmine stage it can be floated off in a staining jar of 50% acetic acid and dehydration with dioxan carried out as above.

Insect salivary gland chromosome smears: The glands are crushed under a cover slip in acetocarmine on a slide coated with dried egg albumen. After 20 minutes the area around the cover slip is flooded with 50% acetic acid and the cover slip floats loose so that it can be removed. The above described dioxan dehydrating procedure is then employed.

Squash preparations: Root tips are fixed in some suitable fixative and the Feulgen technic applied. The stained root tips can either be dehydrated by passing thru 3 changes of dioxan and mounting in dioxan-balsam where they are divided into small longitudinal sections by sharp needles, or they can be put immediately into a mixture of 1 part of 50% acetic acid to 1 part of corn syrup where shredding with needles is carried out. A cover slip is put on and separation of the cells completed by tamping or by applying pressure to the cover. This squash method is useful with anthers which are difficult to smear when in the early prophase stages of meiosis.     

Permanent Preparations from Rapid Cytological Technics

The following rapid but reliable method of making permanent preparations from temporary mounts has proved to be very useful.

Pollen mother-cell smears: Smeared anthers are treated hi the usual way with Belting's acetocarmine, except that the cover slip is left off. When correct differentiation is attained the stain is thoroly washed off with 50% acetic acid and the slide flooded with dioxan. This is followed by 2 changes of dioxan for 2 minutes each. A drop of Canada balsam dissolved in dioxan is added and a cover slip applied. In cases where a cover slip has been used at the acetocarmine stage it can be floated off in a staining jar of 50% acetic acid and dehydration with dioxan carried out as above.

Insect salivary gland chromosome smears: The glands are crushed under a cover slip in acetocarmine on a slide coated with dried egg albumen. After 20 minutes the area around the cover slip is flooded with 50% acetic acid and the cover slip floats loose so that it can be removed. The above described dioxan dehydrating procedure is then employed.

Squash preparations: Root tips are fixed in some suitable fixative and the Feulgen technic applied. The stained root tips can either be dehydrated by passing thru 3 changes of dioxan and mounting in dioxan-balsam where they are divided into small longitudinal sections by sharp needles, or they can be put immediately into a mixture of 1 part of 50% acetic acid to 1 part of corn syrup where shredding with needles is carried out. A cover slip is put on and separation of the cells completed by tamping or by applying pressure to the cover. This squash method is useful with anthers which are difficult to smear when in the early prophase stages of meiosis.     

Differential Staining of Gastric Glandular Cells

Fundus of stomach is fixed in 10% formalin (aqueous), Bouin's fluid or 5% trichloracetic acid (aqueous). It is embedded in paraffin, and 7μ sections are cut, mounted, deparaffinized and passed to 70% alcohol and then stained as follows: Mordant 3 min. in saturated Bismarck brown in 70% alcohol. Rinse in 70% alcohol, pass to distilled water, then overstain (2 hr.) in aniline blue, 0.5% solution in 2.5% acetic acid (aqueous). Precipitate the anilin blue with 0.5 ml. of 0.1% methyl violet solution (aqueous) dropped on die slide. Leave on 2 min. or less. Wash and differentiate in 70% alcohol. (Parietal cells dark blue). Stain 30 min. in a mixture of hematein, 0.10g.; A1C13 cryst., 0.05g.; and 70% alcohol 50 ml., prepared just before use and not filtered. Rinse in 70% alcohol and differentiate with an alcoholic extract of saffron (2 g. saffron pistils in 100 ml. 90% alcohol at 60°C. for 6 hr.) while observing the progress of differentiation microscopically. Dehydrate by dropping a 0.1 % solution of acetic acid in absolute alcohol on the section for 30 sec., followed by pure absolute alcohol, xylene, and covering in balsam.     

Differential Staining of Gastric Glandular Cells

Fundus of stomach is fixed in 10% formalin (aqueous), Bouin's fluid or 5% trichloracetic acid (aqueous). It is embedded in paraffin, and 7μ sections are cut, mounted, deparaffinized and passed to 70% alcohol and then stained as follows: Mordant 3 min. in saturated Bismarck brown in 70% alcohol. Rinse in 70% alcohol, pass to distilled water, then overstain (2 hr.) in aniline blue, 0.5% solution in 2.5% acetic acid (aqueous). Precipitate the anilin blue with 0.5 ml. of 0.1% methyl violet solution (aqueous) dropped on die slide. Leave on 2 min. or less. Wash and differentiate in 70% alcohol. (Parietal cells dark blue). Stain 30 min. in a mixture of hematein, 0.10g.; A1C13 cryst., 0.05g.; and 70% alcohol 50 ml., prepared just before use and not filtered. Rinse in 70% alcohol and differentiate with an alcoholic extract of saffron (2 g. saffron pistils in 100 ml. 90% alcohol at 60°C. for 6 hr.) while observing the progress of differentiation microscopically. Dehydrate by dropping a 0.1 % solution of acetic acid in absolute alcohol on the section for 30 sec., followed by pure absolute alcohol, xylene, and covering in balsam.     

Feulgen's Reaction Applied to Protozoa and Small Worms, Mounted In Toto in Venetian Turpentine

Permanent mounts of certain protozoa and small worms are obtained as follows: kill suspensions of the organisms with Feulgen's fixative (6% HgCl₂ in 2% aqu. acetic acid) for 3 to 24 hours. After pipetting off the fixative, add successively: 70% iodized alcohol; ditto, 30 minutes later; 50%, then 35% alcohol; 2 baths distilled water; normal HCl. Transfer to cold water and heat to 60°C for 4 to 5 minutes or longer. Cool under running water; and wash in distilled water.

Stain 1 to 3 hours in Feulgen's fuchsin sulfurous acid (1 g. of a suitable basic fuchsin, e. g. rosanilin hydrochloride, boiled in 200 cc. water, cooled, and allowed to stand 24 hours after adding 20 cc. normal HCl and 1 g. sodium bisulfite). Pass thru 3 baths of 200 cc. distilled water with 10 cc. normal HCl and 1 g. sodium bisulfite. Transfer to water and then to 35%, 70%, and 95% alcohols successively. Counterstain with fast green FCF, orange G or eosin Y in 95% alcohol. Pass thru two changes of absolute alcohol.

Transfer to 10% Venetian turpentine and place in a dessicator; mount after the turpentine has become concentrated.

If sections instead of total mounts are desired, the material should go from absolute alcohol, thru alcohol-xylol and xylol to paraffin (or preferably paraffin of M. P. 56°C with 3% bees-wax). The paraffin may be added to the material in the test tube, and cooled after the organisms have settled. Then break the tube, trim a block, and cut.     

A Controllable Differential Stain for the Hypophysis by Adaptation of the Mallory Triple Stain

A selective and controllable staining method for the hypophysis has been developed with rat material, using Mallory's triple stain as a basis.

Fix in Zenker neutral formol for 6 hours. Longer fixation is undesirable. Transfer to 30% alcohol plus a few drops of a saturated solution of I₂ in aqueous KI over night. Gradually complete dehydration and clear in cedar oil. Infiltrate with a paraffin mixture (paraffin, rubber-paraffin, bayberry wax and beeswax). Section 3-Sμ. Hydrate to distilled water, placing a few drops of a KI-I₂ solution in the 50% alcohol. Stain in 1% acid fuchsia for 30 minutes. Rinse, and differentiate in a weak NH₄OH solution (one drop 28% NH₄OH to 200 cc. HOH). When differentiation is complete, transfer to a 0.5% phosphomolybdic acid solution for 3 minutes, after first stopping the differentiation with a 0.1% HC1 solution and then rinsing with distilled water. Stain for one hour in a solution of: 1% anilin blue, water soluble, 2% orange 6, and 1% phosphomolybdic acid. Rinse in distilled water plus a few cubic centimeters of the stain. Differentiate in 95% alcohol, transfer to absolute alcohol and clear in a mixture of 30% oil of cedar, 40% oil of thyme, 15% absolute alcohol and 15% xylene. Finally, transfer to xylene and mount.     

Comparative Study of Dehydration

The paraffin method has frequently been criticised because of its hardening and shrinking effect on tissue. The author believes this distortion is due to the dehydration and not to the immersion in melted paraffin. An experimentally controlled series of various tissues was dehydrated in different dehydrating reagents, dioxan, isobutyl alcohol, and ethyl alcohol with chloroform. Except for the dehydration, the tissues were treated identically. In every case, dioxan proved to be a better dehydrating reagent with less shrinkage and brittleness than any of the others. Ethyl alcohol with chloroform produced the greatest degree of distortion.     

Staining Paraffin Sections with Protargol 6. Impregnation and Differentiation of Nerve Fibers in Adrenal Glands of Mammals

A series of experiments with protargol staining of nerve fibers in mammalian adrenal glands has yielded the following procedure: Fix-1-2 days in a mixture of formamide (Eastman Kodak Company) 10 cc, chloral hydrate 5 g., and 50% ethyl alcohol 90 cc. Wash, dehydrate and embed in paraffin. Cut sections about 15 and mount on slides. Remove the paraffin and run down to distilled water. Mordant 1-2 days in a 1% aqueous solution of thallous (or lead) nitrate at 56-60°C. Wash thru several changes of distilled water and impregnate in 1% aqueous protargol (Winthrop Chemical Company) at 37-40°C. for 1 to 2 days. Rinse quickly in distilled water and differentiate 7-15 seconds in a 0.1% aqueous solution of oxalic acid. Rinse thru several changes of distilled water for a total time of 0.5 to 1.0 rain. Reduce 3-5 rain, in Bodian's reducer: hydroquinone 1 g., sodium sulfite 5 g., distilled water 100 cc. Wash in running water 3-5 min. and tone 5-10 min. in a 0.2% gold chloride solution. Wash 0.5 min. or more and reduce in a 2% oxalic acid solution to which has been added strong formalin, 1 cc. per 100. (Caution. This last reduction is critical and over-reduction can spoil an otherwise good stain; 15-30 seconds usually suffices, and the sections should show only the beginning of darkening to a purplish or gray color.) Wash, fix in hypo, wash, dehydrate and cover.     

Delafield's Hematoxylin and Safranin for Staining Meristematic Tissues

The following technic is suggested for staining permanent preparations of meristematic tissues: Prepare and mount the sections by the usual paraffin method. From water, stain them 2-10 minutes in a solution made by adding 2-4 cc. of Delafield's hematoxylin to a Coplin jar full of tap water. As staining is progressive, the sections should be examined from time to time with a microscope. When the cell walls have become a deep purple, transfer the preparations, thru the usual series, to a mixture of xylol-absolute-alcohol in equal parts, and from this to a counterstain made by adding 4-6 cc. of a saturated solution of safranin in absolute alcohol to a Coplin jar full of xylol (75%) with absolute alcohol (25%). This stains the nuclei. Leave the sections in the counterstain at least 2 hours and then rinse them in xylol-absolute-alcohol (1:1) to remove excess safranin. Transfer them to pure xylol and then mount them in neutral balsam.     

Basic Fuchsin and Picro-Indigo Carmine: A Selective Stain for Cells in Mitosis and for Autoradiography

Selective staining of dividing nuclei is accomplished as follows: paraffin sections, after hydration, are stained 15 min in a saturated aqueous solution of basic fuchsin, washed, then stained 1.5 min in an equal-volumes mixture of indigo carmine saturated in 70% alcohol, and saturated aqueous picric acid. Removal of excess dye with 3 changes of 70% alcohol, dehydration, clearing and covering in a resinous medium completes the process. Nuclei of dividing cells are stained red; cytoplasm and interphase nuclei, light green. This method has been used successfully for determining the mitotic activity of skin, kidney, liver and other rabbit and mouse tissues. Tissue sections previously prepared as autoradiographs may be stained by this method to facilitate the determination of radioactive labeling of mitotic cells.