Cytological Methods for Crepis Species

Root tips of Crepis species are fixed in La Cour's “2BE” and dehydrated thru a butyl alcohol series. They are stained in 1% crystal violet for 1 hour, with chromic acid and iodine as pre-and post-staining mordants, respectively, and passed thru dehydrating alcohols containing picric acid and ammonium hydroxide. Differentiation is done in clove oil. The method is rapid; the chromosomes are dark purple; the centromere is not stained; and the cytoplasm is clear. By further controlled destaining the hetero-chromatic segments within the chromosomes may be located.

Pollen mother cells are fixed in acetic alcohol (1:4) and squashed in aceto-carmine. A method is described for making semi-permanent preparations mounted in diaphane.

Pollen grains are mounted in lacto-phenol with acid fuchsin or anilin blue W. S. as the dye.     

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.     

An Easily Controlled Regressive Trichromic Staining Method

Three modifications of Mallory's connective tissue stain are described and some features of the action of picric acid are discussed.

In the first and most critical method the nuclei are stained in an iron hematoxylin and then differentiated in a picric acid solution containing orange G. This not only differentiates the nuclei, but stains all other elements yellow. The section is then washed in running water to remove the yellow color from all tissues except those which are to remain yellow in the final preparation (usually the erythrocytes). The section is next stained in an acid fuchsin mixture and then differentiated until the desired depth and contrast is obtained. Staining in anilin blue follows and this in turn is differentiated to suit. The section is then dehydrated and mounted.

In the second method the nuclei are stained in hemalum (e.g. Harris's) for a short time; the section is then rinsed and immersed in a mixture of picric acid and acid fuchsin and thereafter is differentiated; it is next passed into anilin blue w. s. and then differentiated and mounted as before. This is less critical than method I, but can be applied to large batches of slides at a time.

The third method is a one-solution method. After staining the nuclei in hemalum, the section is immersed in the “Picro-Mallory” solution, differentiated briefly, dehydrated and mounted. This modification, while being the least critical, is most suitable for routine use when the tissues have been fixed in a fluid containing chromate; the other commonly used fixatives, while giving useful results, are not so good.     

Brilliant Cresyl Blue as a Stain for Plant Chromosomes

Methods are proposed for staining plant chromosomes with the dye brilliant cresyl blue, and for making these stained preparations permanent by using polyvinyl alcohol mounting medium.

The stain, which is composed of 2% brilliant cresyl blue in 45% aqueous acetic or propionic acid, is used with fixed material in making smear preparations. The technics for staining are similar to those employed in the aceto-carmine method.

The mounting medium is made by mixing 56% polyvinyl alcohol, which is diluted in water to the consistency of thick molasses, with 22% lactic acid and 22% phenol by volume. The permanent slides are made by floating off the cover slip of the temporary slide in 70% alcohol, then applying the mounting medium and replacing the cover slip.

The chief advantages of the methods described are:

1)The preparation of the stain is rapid and simple. The batch of stain will be good with the first try.

2)The staining procedure in some instances is shorter than when using aceto-carmine.

3)The stain shows a high degree of specificity for nuclear structures and gives better results than aceto-carmine when used on certain plant tissues.

4)A minimum number of cells is lost in making the slides permanent when using polyvinyl alcohol mounting medium as the slide and cover slip are run through only one solution prior to mounting.

5)The mounting medium dries rapidly and this shortens the time required before critical examination of the permanent mounts can be made.     

Staining Methods For Studying Ingredient Distribution In Baked Cakes

A method is described for preparing cake crumb for sectioning and staining. Previous to embedding, the fat was stained and fixed by exposing small blocks of cake to the fumes from a 5%, freshly-prepared, aqueous solution of osmic acid (OsO₄). This was followed by dehydration in ethyl alcohol and tertiary butyl alcohol, removal of air under vacuum and infiltration with paraffin.

Sections were cut 20 and 9Op thick and mounted with water.

Wax was removed by immersion in xylene. The sections were rehydrated in a series of ethyl alcohol dilutions, from concentrated to dilute, then transferred to distilled water.

Protein was then stained pink by immersion of the slides in an acidified 0.04% water solution of eosin Y, or starch was stained blue with a dilute aqueous solution of iodine. Ten grams iodine and 10 g. KI were dissolved in 25 ml. distilled water. This stock solution was diluted for use one to two hundred times.

The relationship between protein and starch was demonstrated by staining the sections with eosin, differentiating in 50% alcohol and staining with iodine.

When slides of cake crumb were prepared in this way, the fat was stained black, the protein bright pink and the starch granules a dark blue.     

The Masson Trichrome Staining Methods in Routine Laboratory Use

A resume of Masson's trichrome staining methods is given, with detailed directions for carrying out all of his procedures. The results obtained thru their use in a routine laboratory are discussed at length, as well as the fact that they also work very well on tissues fixed in ways other than those he prescribes, and stained with chemicals and dyes other than those he uses. The fact is stressed, however, that the closer one adheres to his precepts, the better will be the results.

The stains described include bis hematozylin-phloxine-saffron, his iron-hematozylin-ponceau-anilin-blue, his variants of this stain (of which the light green stain is excellent), his metanil yellow and his modification of the familiar Van Gieson technic. All these stains are based on familiar laboratory methods, improved and rendered trichrome, so that they present no great obstacles in technic.

Of the methods cited, the writer prefers the “light green” procedure. Sections are prestained in Regaud's iron-hematoxylin, followed by a mixture of ponceau de xylidine and acid fuchsin. This is followed by mordanting in phosphomolybdic acid and the sections are finally stained in light green. The results are very precise and pleasing and afford immediate orientation as the connective tissue is green, the nuclei black or dark purple, the cytoplasm of the cells is in varying tones of red. The method may be used after fixation in almost any good medium; altho the results are not as brilliant as those obtained after one of Masson's prescribed fixations, it is believed that they are even then superior to those following the routine hematoxylin-eosin method.     

Permanent Stained Preparations of Thick Blood Films

Fixing thick films in alcoholic solution of dye after the usual staining-and-laking procedure preserves the appearance of parasites and blood elements very similar to that of the usual thick films (not fixed) for the diagnosis of malaria and relapsing fever.

Procedure recommended: Films are stained and laked for 15 minutes in diluted Giemsa—1 to 3 drops of stock solution (0.4 g. in 60 ml. equal parts absolute methyl alcohol and glycerin) per ml. distilled water; rinsed in water and allowed to dry. They are then immersed in, or flooded with, May-Griinwald's stain (0.5% in absolute methyl alcohol) for 30 seconds, rinsed in water and allowed to dry. Solutions of MacNeal's tetrachrome stain in methyl alcohol and glycerin may be substituted for Giemsa and a solution in methyl alcohol may be substituted for May-Griinwald. With slight modification of the procedure, both thick and thin films on the same slide may be stained together.

Films stained and fixed as described, and mounted in Diaphane, have shown no evidence of fading in 3 years.     

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.     

Modifications of Lebowich's Soap-Wax Technic

Lebowich's technic is outlined for simultaneous dehydration and infiltration of tissues by a medium composed of stearic acid, 56° C. paraffin, diethylene glycol, and monoethanolamine. The prices and places where these materials may be purchased are given.

Tissue for sectioning is placed in acetone, C.P., for 1 hour, then put directly into the soap-wax medium at 60° C. under reduced pressure, and finally embedded in new soap-wax.

Modifications include a simplification of the apparatus used by Lebowich. A preserving jar fitted with a rubber stopper serves as a vacuum chamber, and use of an aspirator accomplishes the reduction of pressure. With invertebrate embryos up to 1000 μ diameter no reduction of pressure is needed. Embryos are fixed, washed, placed in acetone, infiltrated in soap-wax, and embedded.

By this soap-wax method the alcohols, xylene, and overnight drying of affixed ribbons are eliminated. Tissue may be fixed, sectioned, stained, and permanently mounted within 6 to 8 hours.     

The Use of Bismarck Brown Y in some new Stain Schedules

The staining quality of Bismarck brown Y may be improved and sterility maintained by adding 5% phenol to a 1% aqueous solution. Use the phenolic Bismarck brown in combination with iron alum hematoxylin except for stripped epidermis in the following procedures:

Stem and Root Schedule: Mordant sections from water in 4% iron alum for 10 minutes. Rinse in distilled water and stain in 0.5% aqueous hematoxylin for 1 minute or until darkly stained. Rinse in distilled water and destain in 2% iron alum until a gray color appears. Rinse thoroly in distilled water and intensify hematoxylin by transferring sections to 0.5% aqueous lithium carbonate until the desired black color appears. Rinse thoroly in distilled water and stain for 1-5 minutes in phenolic Bismarck brown. Rinse in distilled water, dehydrate successively in 30, 50, 70, 95 and 100% alcohol. Clear in methyl salicylate for 5 minutes, then to xylene for 3-5 minutes, and mount in balsam.

Middle Lamellae in Wood: Destain more thoroly in 2% iron alum than for the general stem and root schedule, and intensify in lithium carbonate for a longer period (about 1 hour).

White Potato Tuber Sections: Modify above schedule by reducing time of destaining in 2% iron alum to about 30-60 seconds and intensify hematoxylin until starch grains appear bluish in color. Stain in phenolic Bismarck brown for 1-2 minutes.

Wheat Grain Sections: Fix grain for sectioning when in “dough” stage. Use schedule the same as for potato tuber except for reducing time of staining in phenolic Bismarck brown to about 45 seconds.

Tradescantia zebrina Epidermis: Strip epidermis from leaf while submerged in water. Fix in 100% alcohol 10 minutes, pass thru 95, 70, 50, 30, and 10% alcohol to water. Stain in phenolic Bismarck brown for 10-20 minutes. Dehydrate, clear in methyl salicylate and mount in balsam.     

The Fixation of Tissues Vitally Stained with Trypan Blue

The following fixative is recommended for tissues vitally stained with trypan blue: Chloroform, 2 parts; absolute ethyl alcohol, 2 parts; glacial acetic acid, 1 part; mercuric chloride to the point of saturation.

The tissue should be fixed 1 to 2 hours; transferred to 95% ethyl alcohol for 12 hours; to absolute alcohol for 12 to 24 hours; to a mixture of absolute alcohol and xylol for 1/2 hour, and finally to xylol, before embedding in paraffin. Cedar oil may be used for clearing in the place of xylol; in that case the tissues should be transferred from absolute alcohol to a mixture of absolute alcohol and cedar oil for 24 hours before placing in cedar oil alone.

Various counterstains can be used; Mayer's carmalum is excellent.     

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.     

The Dry Preservation of Fixed Plant Material

A method for the dry-preservation of fixed plant material, root tips and buds, is described. The method seems to be advantageous on long expeditions and when material has to be sent away.

The material is transferred from the fixative to 70% alcohol (3 changes, 1/2 hour in the last). It is dried on blotting-paper. The dried material may be preserved a long time. Material kept dry for 4 1/2 years has proved to be quite satisfactory. Drying has been tried after fixation with CRAF-solutions (Webber and Randolph modifications) and fixatives containing osmic acid (Fleming-Benda and 2BD).

The dry material is swollen by keeping for 2 days in 10% alcohol. It is embedded in paraffin according to the usual method. A satisfactory staining has been obtained after these fixatives using iodine-gentian-violet and Feulgen stainings. In addition to chromosome counts dry material may be used for chromosome morphology studies.

Dried material fixed in aceto-alcohol (1:3) has not turned out to be specially suitable for squash preparations owing to the fragility of the chromosomes. If strong pressure is not applied, satisfactory results may, however, be obtained.     

A Colloidal Silver Method for Nerve Cells and Processes, Neuroglia and Microglia

A method is described whereby nerve cells and processes, neuroglia and microglia may be stained using colloidal silver solutions (argyrol, silvol, 10% and 20%).

Fresh, unfixed brain tissue is stained in bulk in argyrol or silvol, and then dehydrated, embedded in low viscosity nitrocellulose, and sectioned. Before reduction the sections are treated with gold chloride to replace the silver. Sections are reduced in a formalin hydroquinone solution, fixed in sodium thiosulfate, dehydrated, and mounted in euparal. A method is described for removing the nitrocellulose before mounting.

No variation in the method was found to be necessary for the various species tested (rat, guinea pig, rabbit, and dog).     

Suggested Counterstains for Davenport Reduced Silver Preparations of Peripheral Nerves

—Peripheral nerves which have been fixed in a mixture of formaldehyde and acetic acid and stained according to the method of Davenport can be successfully counterstained for demonstration of myelin sheaths and stroma. After mounted sections have been silvered, reduced and toned, the coating of nitrocellulose is removed by passing thru two changes of acetone. Following brief washes in 100,95,85 and 75% alcohols they are stained in an acidified aqueous solution of azo carmine for 30 to 60 minutes. Excess azo carmine is extracted with anilin alcohol followed by acetic alcohol after which the sections are mordanted for 15 to 60 minutes in a 5% aqueous solution of phosphotungstic acid. Without washing they are transferred to a stain mixture of either anilin blue and orange G (acidified) or light green and orange G (acidified) where they remain from 1 to 5 hours. After destaining in 95% alcohol and dehydration in absolute alcohol the sections are mounted in dammar. Result: axons stain black; sheath and fibroblast nuclei, red; myelin sheaths, orange; and connective tissue, blue or green. When the counterstains are applied to ganglia, cytological details of individual cells are demonstrated.     

A Simplified Stain for Hemoglobin in Tissues Or Smears Using Patent Blue

The following technic, based on the patent blue V hemoglobin reaction, is useful for identifying hemoglobin in tissue fixed in neutral formaldehyde solution and embedded in paraffin:

Stain the deparaffinized, hydrated sections 3 to 5 minutes in the working reagent, prepared by adding 2 ml. of glacial acetic acid and 1 ml. of 3% hydrogen peroxide to 10 ml. of the filtered stock solution (1 g. patent blue, 10 g. zinc powder, and 2 ml. glacial acetic acid). Counterstain 30 to 60 seconds in 1:1000 safranin solution in 1% acetic acid, rinse, dehydrate with alcohols, clear in xylene and mount in clarite. Total time required, 37 minutes.

Blood and tissue and smears may be stained, following fixation in methyl alcohol, by applying the working reagent as above.     

Mallory's Connective Tissue Stain Following Hematoxylin

The following combination of hematoxylin with Mallory's connective tissue stain is useful in bringing out nuclei as well as in differentiating tissue:

Slightly overstain in Mayer's hematoxylin (50 g. potassium alum and 0.2 g. sodium iodate added to 1 liter 0.1% aqueous hematoxylin). Wash; and stain 30 seconds to 1 minute in 0.04% aqueous acid fuchsin-Stain 4 minutes in: 0.5 g. anilin blue and 2 g. orange G dissolved hi 100 cc. of 1% aqueous phosphomolybdic acid. Pass thru 95% alcohol to absolute; clear in xylol and mount in balsam.     

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 Staining Certain Bacteria and Antherozoids

A modification of Loeffler's method for staining the flagella of bacteria was employed in staining large forms of bacteria and antherozoids. The bacteria or the antherozoids are killed and fixed in a drop of water on a slide and set aside to dry, before the next step is undertaken. The slide is treated for a period of time, varying from about ten minutes to several hours, in a practically saturated solution of tannic acid. After the slide is thoroly rinsed in water, it is stained with either a single stain or a combination of stains. The slide is then dehydrated with absolute alcohol, cleared, with clove oil, and completed in the usual manner.

The body of the bacterium and that of the antherozoid are well differentiated and the cilia are distinctly brought out by means of the method herein described.

The technic is of especial value in staining the antherozoids of mosses, liverworts, and ferns.