Book - A text-book of histology arranged upon an embryological basis (1913) 2-1

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Lewis FT. and Stöhr P. A Text-book of Histology Arranged upon an Embryological Basis. (1913) P. Blakiston’s Son and Co., 539 pp., 495 figs.

Histology with Embryological Basis (1913):   Part I. 1.1. Cytology | 1.2. General Histology | 1.3. Special Histology Part II. 2.1. The Preparation of Microscopical Specimens | 2.2. The Examination of Microscopical Specimens

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Part II. Microscopical Technique

I. The Preparation Of Microscopical Specimens

Revised By Lawson G. Lowrey.

The methods of fundamental importance, which are likely to be employed by students who are beginning their histological studies, are here given. Further information may be obtained from "The Micro tomist's Vade Mecum" by A. B. Lee (Blakiston, Philadelphia) and from Mallory and Wright's "Pathological Technique" (Saunders, Philadelphia). The former deals with the subject from the point of view of general biology; the latter is particularly adapted to the needs of medical students.

FRESH TISSUES.

Certain tissues may be studied advantageously in a fresh condition. They are simply spread on a clean glass slide, covered and examined. Desquamated epithelial cells, spermatozoa, blood, and other fluids containing cells, may be treated in this way. But structures such as muscles, tendons, nerves, connective tissue, etc., must first be "teased" that is, torn into very small fragments or spread into a thin layer with a pair of fine needles.

The "parenchymatous" organs, or other structures which cannot be investigated satisfactorily by the above methods, must be sectioned or macerated. The old methods of making free-hand sections of the object held between pieces of pith, or of making sections with a double bladed knife, have been superseded in most laboratories by the freezing method. This method is often serviceable in histology, and is indispensable in the rapid diagnosis of pathological conditions.

Blocks of tissue not over 5 mm. thick are moistened with water, placed on the carrier of a special form of microtome and frozen by a jet of carbon dioxide proceeding from a tank of the compressed gas. Sections 10 to 15 // thick may be chiselled from the block of tissue and unrolled by transferring to a dish of 0.6 per cent, sodium chloride solution. They are floated on a slide, covered and examined.

487

488 HISTOLOGY

Sections or teased preparations must be kept moist during examination. In order to avoid distortion, they are not mounted in water, but in socalled indifferent fluids, such as the lymph, aqueous humor, serous fluids, amniotic fluid, etc. Of the artificial indifferent media, a 0.6 per cent, solution of sodium chloride in distilled water has been found to cause less distortion than the stronger fluids formerly recommended.

Ringer's Solution. An indifferent fluid which is perhaps more satisfactory than the 0.6 per cent, salt solution is a modification of Ringer's solution adapted to the tissues of warm-blooded animals. It is to be made in large quantities.

Sodium chloride 90. o

Potassium chloride 4.2

Calcium chloride (anhydrous) 2.4

Potassium bicarbonate 2.0

Distilled water 10,000.0

Examination of fresh tissues reveals but little of the fine details of structure. Since the indices of refraction of the different tissue elements have much the same value, outlines are usually dim and there is very little optical differentiation. The method of handling is prone to produce distortion and with many tissues and organs it is difficult to separate their constituent elements. It is generally necessary to employ more complex methods of treatment to gain an adequate idea of the histological details.

One of the simplest methods is to add one or two drops of i to 5 per cent, acetic acid solution to the fresh preparation. The nuclei then appear more distinctly. Albuminous granules are dissolved, but fat and myelin are not affected. The white fibers of connective tissue swell and disintegrate, leaving the elastic fibers unaffected.

Nuclei may be rendered distinct by allowing a few drops of stain to act upon the tissue for a few minutes. A i per cent, aqueous solution of methylene blue, or a i per cent, solution of methyl green in 20 per cent, alcohol, or the haematoxylin solutions, may be used.

ISOLATION.

Some tissues cannot properly be separated into their elements in the fresh condition, but may be shaken or teased apart after preliminary treatment. The reagents employed in maceration have the property of softening or removing certain constituents of the tissues, at the same time fixing or hardening other elements. Usually the intercellular portions are softened or removed, while the cellular elements undergo fixation.

Ranvier's Alcohol. This is a mixture of one volume of 95 per cent, alcohol and 2 volumes of distilled water. The cells of small pieces of epithelium (5-10 mm. square) are separable in 24 to 48 hours. They are examined in the same fluid, or washed in water and examined in glycerin.

MACERATING FLUIDS 489

Nitric Acid and Potassium Chlorate. About 5 gm. of potassium chlorate are dissolved in 20 c.c. of the acid. Muscle cells are separable in one to six hours. Wash thoroughly in water and examine in water or glycerin.

Potassium Hydrate. Muscle cells may be teased apart after immersion for about an hour in a 35 per cent, aqueous solution. They may be examined in the same solution or transferred to a saturated aqueous solution of potassium acetate, which prevents further maceration. The solution of potassium hydrate may also be used for isolating epithelial cells.

Concentrated Sulphuric Acid. The elements of the epidermis, hair and nails may be separated after immersion in this fluid. They should be thoroughly washed in water.

PERMANENT PREPARATIONS.

None of the methods described above yield much information respecting the finer structure of tissues and organs, nor do they yield permanent preparations. For ease of reference, the various steps in the production of a permanent preparation have been grouped under the following five headings.

1. Fixation. Under this heading are given formulae for the best fixing fluids, with directions for their use and for the subsequent handling of the tissue until it is placed in 80 per cent, alcohol, in which tissues may be kept for a considerable time.

2. Imbedding. This includes the various steps for preparing the tissues to be sectioned in paraffin or celloidin, starting from 80 per cent, alcohol.

3. Cutting and handling sections. Brief directions are given for cutting sections and handling them, until they are ready for staining.

4. Staining. Formulae and directions for the use of stains, and the after treatment until the preparation is in the appropriate clearing fluid.

5. Clearing and mounting. The choice of a clearing agent for paraffin and celloidin sections rs discussed, together with the methods and media for mounting.

Since each of the fixing, imbedding and staining methods is considered as a unit, each starting where the previous step ends, the student can easily prepare specimens according to any desired possible combination by referring to the directions for the selected fixative, imbedding method, and stain.

i. Fixation.

A good fixative should penetrate and kill tissues quickly; preserve the tissue elements, particularly the nuclei, in the condition in which they are

4QO HISTOLOGY

found at the moment of its action; render structures insoluble, and harden them so that they will not be altered by the various after-steps; and give a certain degree of optical differentiation.

No single compound has yet been found which successfully fulfills all of these conditions, nor are any of the recommended fixatives adequate in all cases or for all special studies. Only the fluids commonly employed, which have proven most useful, are here given.

Small pieces of tissue, preferably less than i cm. in thickness, should be dropped into a considerable amount of fluid. The tissue should be handled as little as possible, in order that delicate structures may not be destroyed. For example, contact between the fingers and the peritoneum is sufficient to destroy the thin epithelium.

In order to insure uniform action of the fixing fluid, it is often advisable to place a little absorbent cotton in the bottom of the vessel. Frequent gentle mechanical agitation will serve the same end. Tubular organs should be washed out, or cut open and their contents and any adherent blood washed away, with salt solution. Membranes may be kept flat and smooth by tying them across the end of a short tube or detached bottle neck.

Alcohol. Small or thin pieces of tissue are supported on a little absorbent cotton in absolute alcohol, for 12 to 24 hours, changing after 3 or 4 hours. Large pieces are fixed by successive immersion in 70 per cent., 80 per cent., and 95 per cent, alcohol for 24 hours each.

Alcohol is a valuable dehydrating and hardening agent, but its fixing qualities are inferior, so that it is rarely used alone as a fixative. Small embryos or blocks of tissue obtained in an emergency should be preserved in 10 per cent, formalin, rather than in alcohol.

Bouin's Fluid.

Picric acid, saturated aqueous solution 75 '

Formalin 20

Glacial acetic acid 5

This fluid is particularly recommended for the fixation of embryos, for which it is unexcelled. Small embryos are fixed in 4 to 6 hours. Larger objects may be fixed 24 to 48 hours or longer. For washing out the fixing fluid, alcohol, first 70 per cent., then 80 per cent., should be employed. Renew the alcohol as often as discolored.

Carney's Mixtures.

No. i Absolute alcohol 6

Chloroform 3

Glacial acetic acid i

This is a very rapid fixative, even large pieces being fixed in $ to X hour. Wash in absolute alcohol until the odor of acetic acid is lost, changing every 12 hours, and imbed; or grade through 95 per cent, to 80 per cent, alcohol.

FIXING FLUIDS 49 1

No. 2. Saturate mixture No. i with mercuric bichloride (about 20 parts). This is the most rapid and penetrating fixative known, and it affords a very delicate cytological fixation. Immersion for 30 minutes to i hour is sufficient even for the larger pieces. Subsequent treatment as with No. i, except that the crystals of sublimate must be removed from the tissue, either by placing the block in 80 per cent, alcohol and iodine (see Zenker's fluid) ; or after the block has been cut, by treating the sections with iodine (see p. 497).

Flemming's Fluid.

Osmic acid, i % aqueous solution 10

Chromic acid, i % aqueous solution 25

Glacial acetic acid, i % aq. solution 10

Distilled water 55

This solution should be mixed only at the time of using. Only very thin pieces (not over 2 mm. thick) should be used. Fix for 24 hours or longer (sometimes even for weeks). Wash in running water 24 hours. Pass through 50 per cent., 70 per cent. (12 hours in each), to 80 per cent, alcohol.

Formaldehyde. The gas is soluble in water to the extent of 40 per cent., and solutions of this strength are obtainable under the trade names of formalin, formol, and formalose.

For fixing tissues, 10 c.c. of the commercial product are added to 90 c.c. of water. It penetrates very quickly, but specimens may be left in it for a considerable time without apparent harm. Ordinary blocks are sufficiently fixed in from 12 to 24 hours. Transfer directly to 80 per cent, alcohol.

Histologically, its chief use is for the preservation of nervous tissue, the fixation of tissue to be cut with the freezing microtome, and the preservation of embryos. Small human embryos obtained by practitioners should be put at once into 10 per cent, formalin and forwarded to an embryological laboratory.

March!' s Fluid.

Potassium bichromate 2.5

Sulphate of sodium i . o

Water 100 . o

Osmic acid, i% aqueous solution 50.0

Small pieces are fixed for 5 to 8 days in the dark. Wash 24 hours in running water; 50 per cent, and 70 per cent, alcohol (24 hours each); 80 per cent, alcohol. Used for demonstrating degenerated nerve fibers and in making damar mounts of fat and myelin, since the osmium reduced by fat is insoluble in alcohol. Sections must not be treated with xylol, but chloroform should be used instead.

Orth's Fluid.

Potassium bichromate 25

Sodium sulphate 10

Water. . . . 1000

492 HISTOLOGY

At the time of using mix 10 c.c. of formalin with 90 c.c. of the above solution (which is known as Miiller's fluid). Small pieces are fixed in about 48 hours. Wash in running water for 12 to 24 hours. Then 50 per cent, alcohol and 70 per cent, alcohol, 12 to 24 hours each; 80 per cent, alcohol. This is useful as a fixative for the central nervous system, and as a general fixative.

Zenker's Fluid. This is kept in the form of the following stock solution, in preparing which the water is heated and the ingredients are stirred with a glass rod (metal instruments must not be put into this fluid) .

Potassium bichromate 25

Sodium sulphate 10

Mercuric bichloride 50

Water 1000

At the time of using, add 5 c.c. of glacial acetic acid to 95 c.c. of the above solution. The tissues, which float for a short time, are fixed for 6 to 24 hours, after which they are washed in running water 12 to 24 hours. Then they are transferred to 50 per cent, alcohol for 12 to 24 hours; 70 per cent, alcohol, 12 to 24 hours; 80 per cent, alcohol.

Corrosive sublimate forms crystalline deposits in the tissues, and these must be removed before the preparation is stained. They may be removed by adding enough tincture of iodine to give a port-wine color to the 70 per cent, and 80 per cent, alcohols in which the block of tissue is immersed. More iodine is added as the solution becomes colorless (or nearly so) and the treatment must be continued until the color no longer changes. The tissues are then to be placed in fresh 80 per cent., renewed two or three times in order to remove completely the mercuric iodide. The crystals of sublimate may be removed after the tissue has been sectioned, as described on p. 497.

Zenker's fluid is an excellent fixative, which penetrates easily and does not decrease the staining qualities. It is probably the best "general fixative."

DECALCIFICATION.

Specimens which contain bone or calcareous material cannot be sectioned until they have been decalcified. The tissues are fixed, according to the directions given above, in Zenker's fluid, Orth's fluid, or formaldehyde, and hardened. After several days in 80 per cent, alcohol, they are put into a considerable quantity of 3 to 5 per cent, aqueous solution of nitric acid. This should be renewed at intervals for 3 or 4 days, until the bone can be penetrated easily with a needle. Wash in running water for a day, and return to 80 per cent, alcohol. Imbed in celloidin.

Phloroglucin is sometimes added to the decalcifying fluid to protect the tissue. The following solution has been recommended. It is to be used in the same manner as the aqueous solution of nitric acid.

DECALCIFYING FLUIDS 493

Phloroglucin i

Nitric acid 5

Alcohol, 95% 70

Water 30

The addition of i or 2 per cent, of nitric acid to the 80 per cent, alcohol will decalcify small embryos. The specimen should then be thoroughly washed in fresh 80 per cent., in order to remove the acid.

2. Imbedding.

Most of the fixatives employed are in aqueous solution. After fixation and the removal of the fixative by washing in water or alcohol, as directed, the specimen must not be left in water, but must be dehydrated. Dehydration has a double purpose: (i) to remove the water, which especially favors post-mortem decomposition, and (2) to prepare the tissue for infiltration with the imbedding substance or, in the case of objects to be mounted whole, for infiltration with the mounting medium.

All the fixation methods given above end with placing the block of tissue in 80 per cent, alcohol. Here they may be left until wanted, although immersion for a considerable time causes a gradual loss in staining qualities. Stronger alcohol causes an overhardening, while maceration may occur in weaker alcohols.

Dehydration is accomplished by immersing the specimen in gradually increasing strengths of alcohol. Those commonly employed are 50 per cent., 70 per cent., 80 per cent., 95 per cent, and absolute. The lower grades may be prepared from the ordinary barrel alcohol, of about 95 per cent, strength, as follows:

80 per cent. 425 c.c. 95 per cent, alcohol mixed with 75 c.c. distilled water 70 per cent 370 c.c. " 130 c.c.

50 per cent. 265 c.c. " 235 cc.

The specimen is left in each grade long enough to be saturated. The time required varies from 3 to 24 hours. Objects of average size require about 6 to 1 2 hours. Prolonged immersion in 95 per cent, or absolute is very injurious to the tissues.

In imbedding, the tissue is surrounded and infiltrated with a firm substance which can be cut into thin sections, supporting and holding firm the fragile tissue. Celloidin, which is solid upon the evaporation of its solvent, and paraffin, which is solid at ordinary temperatures, are the substances used, each having its particular advantages.

Paraffin Imbedding. Specimens cannot be passed directly from alcohol to paraffin, since alcohol dissolves only a very little paraffin and the specimen would not be thoroughly infiltrated. So the specimen must first be passed through some fluid which mixes with absolute alcohol and will dissolve paraffin. Of a host of reagents possessing this property, chloroform is recommended for general use.

After thorough dehydration (12-24 hours in absolute alcohol), the

494 HISTOLOGY

specimen is passed from absolute to a mixture of equal parts of absolute and chloroform for 2 to 6 hours, and then to pure chloroform for an equal length of time. It is then transferred to a saturated solution of paraffin in chloroform, kept warm by placing on top of the paraffin bath, for 2 to 4 hours, and is then put into melted filtered paraffin.

The melting point of the paraffin used varies with the temperature in which it is to be cut. During the winter, paraffin melting at 5o-52 C. should be used, while during the summer paraffin with a melting point of 56-58 is best. Harder paraffin is required for thin than for thick sections. The melted paraffin should be kept in a paraffin bath or thermostat maintained at a temperature but slightly higher than the melting point of the paraffin.

The specimen should be left in the melted paraffin for the shortest time which will allow thorough infiltration, as heat is very injurious to the tissue. For average specimens, 3 hours is sufficient. Transfer to fresh paraffin at the end of i| or 2 hours. At the end of the full time, the specimen is to be imbedded.

The imbedding frame consists of a glass plate and two L-shaped pieces of metal. By sliding the latter back and forth on one another, the size of the enclosed space or box may be varied. Before using the frame, the inner surfaces of the metal pieces and that part of the glass plate on which they rest are rubbed with glycerin. It should form a thin film over the surfaces, but not accumulate in drops. Melted paraffin is poured into the box and the specimen is transferred to it with a spatula. The specimen sinks to the bottom, and may be arranged in any desired position by means of needles warm enough to prevent the paraffin solidifying over their surfaces. The paraffin must be quickly cooled by lowering the frame into a basin of cold water so that the water comes up on the sides of the metal pieces. As soon as a resistant film has formed over the surface of the paraffin, the entire frame may be submerged, and in a few minutes the glass plate and metal pieces may be detached from the solid paraffin. The block may be sectioned as soon as it is thoroughly cooled.

When a number of specimens are to be imbedded; a flat dish of suitable size may be used. After a thin layer of glycerin has been coated over the interior, the dish is filled with a sufficient quantity of melted paraffin and the blocks are put into position. The mass is cooled and removed as before, and the large mass is cut into smaller parts, each containing a specimen.

One or several specimens may be imbedded in paper boxes of suitable size. The tabs at the ends may be labelled and the specimens kept in the boxes until wanted; otherwise labels may be scratched in the paraffin with needles.

PARAFFIN IMBEDDING 495

Paraffin imbedding is to be chosen when very thin sections or serial sections are desired. Material imbedded in paraffin may be kept for years without any apparent deterioration.

Celloidin Imbedding. Thick celloidin is prepared by dissolving 30 gm. of Schering's granular celloidin in 300 c.c. of a mixture of equal parts of ether and absolute alcohol. It has a thick syrupy consistency, and becomes constantly denser by evaporation of the solvent. It should be kept in a tightly closed preserve jar. Thin celloidin is prepared by mixing equal volumes of the thick celloidin and the absolute and ether mixture.

The hardened and dehydrated block of tissue, trimmed to the size and shape desired, is transferred from absolute alcohol to a mixture of equal parts of ether and absolute for 24 hours. From this it is transferred to thin celloidin, in which it remains from 24 hours to a week or longer, and then to thick celloidin for the same length of time. The success of the process depends largely upon the thorough infiltration of the tissue with the celloidin. The time required in the celloidin varies with the penetrability of the tissue and the size of the piece.

After remaining for a sufficient length of time in the thick celloidin, the tissue is taken out with a mass of adherent celloidin and is pressed gently against the roughened surface of a block of vulcanized fiber. As soon as a film has formed upon the surface, the block and attached specimen are dropped into 80 per cent, alcohol, in which the mass becomes firm. It is ready for sectioning in about 6 hours.

In case it is desired to secure sections through the entire thickness of the specimen, the following method is recommended. A sufficient quantity of thick celloidin is poured into a flat dish (or paper box) and the specimen is put into it. The entire mass is hardened as before and then a block of celloidin containing the specimen is cut out. This is trimmed to leave only a thin rim around the specimen. The block is placed for a few moments in the ether-absolute mixture, and then dipped in thick celloidin and pressed against the surface of a fiber block, which has also been dipped in the ether-absolute mixture and in thick celloidin. The mass is allowed to harden somewhat, and then is placed in 80 per cent, alcohol.

The imbedded specimen is kept in 80 per cent, alcohol until wanted for sectioning. Celloidin imbedding is recommended for large objects, or for those from which very thin sections are unnecessary.

Rf SUME" OF IMBEDDING METHODS.

Assuming that the tissues have been fixed and carried into 80 per cent, alcohol, the steps in imbedding are as follows:

496 HISTOLOGY

Paraffin Celloidin

95 per cent alcohol 12-24 hr. 95 per cent, alcohol 12-24 hr.

Absolute 1 2-24 hr. Absolute 1224 hr.

Absolute and chloroform, equal parts. 2 6 hr. Absolute and ether, equal

Chloroform 2- 6 hr. parts 12-24 hr.

Chloroform saturated with paraffin. 2- 4 hr. Thin celloidin 24 hr. to a week

Melted paraffin 2- 4 hr. Thick celloidin 24 hr. to a week

Imbed in fresh melted paraffin and cool quickly. Mount on fiber block; harden and preserve in

80 per cent, alcohol.

3. Cutting and Handling Sections.

Paraffin Sections. Two kinds of microtomes are in general use for sectioning objects imbedded in paraffin. In one form, the "precision microtome," the knife is horizontally placed and the object is moved backward and forward on a carrier. In the rotary microtome, the knife is vertically placed and the object is moved up and down, being cut on the down stroke. In both forms, the knife edge is at right angles to the carrier and the object.

For sectioning with the precision microtome, the object is mounted on a fiber block which is then clamped in the microtome; with the rotary form, it is mounted on a special metal disc. Before attaching the imbedded object, superfluous paraffin is cut away, leaving the tissue rising from a broad base and completely surrounded by a thin layer of paraffin. The block should be trimmed so as to give a rectangular or square surface to be cut, and there should be a considerable layer of paraffin between the object and the block or disc to which it is to be attached. The base is placed upon a heated spatula which rests upon the fiber block. When the paraffin is somewhat melted, the spatula is withdrawn and the base is pressed down upon the block, to which it adheres when the paraffin solidifies. In mounting upon the metal disc, the disc is heated, the block pressed \ipon it and the whole quickly cooled by immersing in water.

If the paraffin on each side of the object is trimmed parallel with the knife edge, the successive sections adhere to one another, forming ribbons. As they are taken from the knife, the ribbons are laid in a shallow box. By placing them in order, they may later be attached to the slide in perfect series, one after the other. The first one cut is attached to the upper left hand corner, and the others follow like lines on a printed page. Sections mounted in this way are called serial sections. The sections should be from 5 to 10 p in thickness, but under favorable conditions thinner sections may be secured.

Before they can be stained, paraffin sections must be attached to the slide and the paraffin must be removed. To attach them to a slide, a mixture of equal parts of white of egg and glycerin is used. The white of egg is thoroughly stirred and filtered. An equal volume of glycerin is added, the two thoroughly mixed and a small lump of camphor added as

PARAFFIN SECTIONS 497

a preservative. The mixture is kept in a glass-capped bottle, with a glass rod for a dropper.

A drop is placed upon a thoroughly clean slide and rubbed evenly with the ringer (freed from oil) over all the area upon which sections may be placed. It should be free from bubbles and should make a very thin layer, just thick enough to allow the finger to glide easily over the surface of the slide. A few drops of water are placed upon it, forming a layer over the albumen deep enough to float the paraffin sections, strips of which are placed upon the water. The shiny side of the ribbon should rest upon the water. The slide is then held for a moment over the flame of an alcohol lamp so that the water is heated. Repeat until the sections become perfectly smooth and flat, but the paraffin must not be melted. The water should not come in contact with the fingers holding the slide. If the albumen layer ends abruptly before reaching the border of the slide, the water will not so readily spread beyond it. After the flattening process, the water is cautiously drained off by a moist sponge held at the corner of the slide. The sections settle down upon the albumen and may be arranged in straight lines with needles applied to the paraffin, but not to the tissues of the sections. The slide is then held vertically in contact with filter paper to drain off any water which may remain, and the portions of the slide which are free from sections are wiped off with a cloth free from lint. The slide is next placed in a drying oven which is not warm enough to melt the paraffin. It is well to let the slides remain over night, but a few hours may be sufficient to dry them thoroughly.

In preparing large numbers of slides, each bearing only one or two sections, fragments of the ribbon containing the desired number of sections are floated in a basin of water warm enough to flatten but not to melt them. Slides rubbed with albumen are dipped into the water beneath the sections, which are held in place with a needle. The slides are drained and dried in the usual way, care being taken to have the sections in the center of the slide. Or the ribbons may be floated on warm water and cut into fragments with a heated knife, proceeding then as before.

To remove the paraffin, the slides are immersed in xylol for about 5 minutes. The slide is then transferred in turn to a mixture of equal parts of xylol and absolute alcohol, then through absolute, 95 per cent., 80 per cent., 70 per cent, and 50 per cent, alcohols, remaining about i minute in each, to water. In case the stain is in alcoholic solution, the transfers may be stopped at that grade of alcohol which corresponds to the solvent of the stain.

In case the sectioned tissue was fixed in a fluid containing corrosive sublimate and has not previously been treated with iodine for the removal of mercurial deposits, enough tincture of iodine to give a port-wine color

may be added to the 80 per cent, alcohol. The slide is immersed in this

32

498 HISTOLOGY

for about a minute and then washed in fresh 80 per cent, to remove the iodine.

All of the reagents and stains to be used for paraffin sections may be kept in a series of tube-like vials, in which the slides may be placed in pairs back to back, being transferred from one vial to another. The vials are kept tightly corked, and the reagents can be used for some time before they must be renewed.

In handling a large number of slides, grooved rectangular boxes are useful. Each reagent is allowed to act, poured out and another substituted.

Celloidin Sections. Objects imbedded in celloidin are cut with either a sliding or a precision microtome, the knife edge meeting the block obliquely. The block and knife are kept wet with 80 per cent, alcohol. Sections are cut 10 to 15 n in thickness, and are transferred, by means of a camel' s-hair brush wet with alcohol, to a dish of 80 per cent, alcohol, in which they remain until wanted for staining.

Celloidin sections are stained in a series of small, shallow staining dishes. The sections are taken from 80 per cent, alcohol and transferred through graded alcohols to water or the solvent of the stain. If deposits of corrosive sublimate are present and were not removed before imbedding, the sections should be treated as directed for paraffin sections. The sections are transferred from dish to dish with bent metal or glass needles. Celloidin sections are not treated with absolute alcohol, since the celloidin would be softened.

The handling of large numbers of celloidin sections is facilitated if they are placed in a perforated cup which fits into another ordinary cup. The ordinary cups contain the various reagents and the sections are transferred from one to another in the perforated cups. The latter may be obtained as "Hobb's Tea Inf users," and lemonade cups are of proper size to receive them.

Wright's Method for Frozen Sections. This method gives permanent preparations which are adequate for most routine purposes in histological examination, and saves much time, labor, skill and expense. The success of the method depends to a considerable extent upon the frozen sections being as thin as good celloidin sections. Special automatic microtomes may now be purchased, or the older form in which the sections are "chiselled" from the block will give good results if properly used.

The tissues are fixed in 10 per cent, formalin for 12 to 24 hours or longer. The piece is then trimmed so that it will present a thickness to be frozen of not over 5 mm. The other dimensions of the block may be as large as the freezing box of the microtome will permit. The block is rinsed in water, placed on the freezing box with a few drops of water beneath it; frozen and cut into sections, which should not be over 10 or 15 p- in thickness.

FROZEN SECTIONS 499

The sections are floated into water, in which they unroll. Select a good section and spread it smoothly on a slide which has been coated with a thin, even layer of albumen-glycerin. Superfluous water is drained off and the section pressed upon the slide with a piece of smooth blotting paper, by exerting an even but not great pressure with the ball of the thumb. The section will adhere to the slide.

Now quickly cover the section with a small quantity of 95 per cent, alcohol, followed in a few seconds by absolute. Pour from a drop bottle quickly and evenly over the section and adjacent surface of the slide a very thin solution of celloidin dissolved in equal parts of ether and absolute alcohol. Drain off immediately, blow the breath once or twice on the surface of the section and immerse the slide at once in water for a few seconds. The thin film of celloidin thus formed fastens the section to the slide. The solution of celloidin should be almost watery in consistence, and so thin that it will form drops readily without stringing. If it is too thin, it will not hold the section on the slide, while if it is too thick, the layer on the slide will become white when it is immersed in water. The film of celloidin should be so thin as to be almost invisible.

The section may now be stained by any of the usual methods applied to sections affixed to the slide. The thin layer of celloidin offers no obstruction to the staining. After staining, the section is dehydrated by covering with 95 per cent, alcohol for a few seconds. Absolute is now poured on and allowed to remain for a few seconds. This removes most of the celloidin, but, unless the action is unduly prolonged, the section will not be loosened. Clear in xylol.

The microtomes and knives mentioned in this section are described and directions for their use are given in Mallory and Wright's "Pathological Technique." Their use, however, is seldom learned except by personal demonstration in the laboratory.

4. Staining.

The purpose of staining is to differentiate the tissue elements. The staining of tissues is in a measure a micro-chemical color reaction, the differential staining being due to the fact that certain elements take up more of the stain than others.

Stains used in microscopic work may be divided into two general classes according to their chemical properties (i) basic stains, which show especial affinity for the nuclei of cells and are called nuclear stains, and (2) acid stains, which affect the cytoplasm more readily and are called cytoplasmic stains. Certain so-called selective stains (either acid or basic) affect one tissue element especially, or even exclusively. Preparations may therefore be stained with several dyes, each affecting certain tissue

500 HISTOLOGY

elements only. Certain stains may be applied to the tissue before it is imbedded and sectioned. They are seldom used except in the preparation of embryos.

GENERAL STAINS.

THE STAINING OF SECTIONS.

Haematoxylin and Eosin. Haematoxylin is a basic dye obtained from logwood, which stains nuclear structures blue. Eosin is an acid anilin dye which stains cytoplasm red. (It is recommended that all anilin dyes used in histological work be those prepared by Griibler in Germany.)

There are many formulae for the preparation of haematoxylin solutions, of which the two following are especially useful.

Alum haematoxylin.

Haematoxylin crystals i gm.

Saturated aqueous solution of ammonia alum 100 c.c.

Water 300 c.c.

Dissolve the haematoxylin in a little water with the aid of heat, and add to the remainder of the solution. Put the mixture in a bottle and add a small lump of camphor or thymol to prevent the growth of mould. Stopper the bottle with a loose plug of cotton and set in the light for about 10 days to ripen. It changes to a deep blue color during this process of oxidation, after which it is ready for use and is kept tightly stoppered. It deteriorates and must be renewed after a few months. The solution may be ripened immediately by the addition of 2 c.c. of hydrogen peroxide solution, neutralized by a crystal of sodium chloride.

For use after Zenker fixation, the water in the above formula may be omitted.

Delafield's Haematoxylin.

Hsematoxylin crystals 4 gm.

Alcohol, 95 per cent 25 c.c.

Sat. aq. sol. of ammonia alum 400 c.c.

The haematoxylin is dissolved in the alcohol and added to the alum solution. This is exposed to the light in an open vessel to ripen for about 4 days and then is filtered. To the filtrate is added:

Methyl alcohol (or 95%) 100 c.c.

Glycerin 100 c.c.

This mixture is exposed to the light in a cotton-plugged bottle for about a week, after which it is again filtered and tightly stoppered. The solution keeps for a considerable time. It may be used in this strength, but preferably it is diluted with one or two volumes of water.

Eosin is sold in two forms, one soluble in water, the other in alcohol. In connection with haematoxylin, a T V to i per cent, aqueous solu

HjEMATOXYLIN AND EOSIN 501

tion may be used, but it is preferable to use a solution made by adding to 95 per cent, alcohol enough of a stock solution, consisting of 5 gm. of eosin dissolved in 300 c.c. of 50 per cent, alcohol, to give a deep yellowishred color to the alcohol.

Sections are transferred from water to the haematoxylin solution for about 5 minutes. They are then washed in several changes of tap water until the sections are deep blue in color. If this change does not occur rapidly, a few drops of ammonium hydrate may be added to the water They are then examined with the microscope. If the sections are overstained (a condition recognizable in the staining of the cytoplasm as well as the nuclei) the sections are washed in a 0.5 per cent, solution of hydrochloric acid in 70 per cent, alcohol until the section is reddish-brown, usually about i minute. Wash in water made alkaline with a few drops of ammonia and re-examine. If the nuclei are not well stained, the sections are returned to the haematoxylin solution for 5 to 10 minutes longer, after which they are examined as before.

When the stain is sharply limited to the nuclei and is of satisfactory depth, the sections are washed for 15 to 30 minutes in tap water and then are passed through 50 per cent., 70 per cent., 80 per cent, and 95 per cent, alcohol, remaining about i minute in each. Stain in the alcoholic solution of eosin for i to 5 minutes. Wash in 95 per cent, until red clouds no longer leave the section.

Paraffin sections are then passed through absolute; absolute and xylol (equal parts) ; and xylol, in which they remain for about 5 minutes, and are then ready for mounting. Celloidin sections are transferred from 95 per cent, to oil of origanum for about 5 minutes, before mounting.

Eosin and Methylene Blue. This method is highly recommended, especially for tissues fixed in Zenker's fluid and sectioned in paraffin.

Stain the sections for 20 minutes or longer in a 5 or 10 per cent, aqueous solution of eosin. The tissue must be overstained, as the eosin is partially extracted in the subsequent treatment. Wash out the excess of the stain in water.

Stain for 10 to 15 minutes in Unna's alkaline methylene blue (i gm. of methylene blue and i gm. of potassium carbonate dissolved in 100 c.c. of water) diluted i : 3 or i : 5 with water. Wash in water. Differentiate and dehydrate in 95 per cent, alcohol, keeping the section in constant motion so that the decolorization is uniform. When the pink color returns to the section and when, as seen under the microscope, the blue is limited to the nuclei, the section is quickly washed in absolute, passed through absolute and xylol, and put in xylol for about 5 minutes.

Heidenhain's Iron Haematoxylin. The best results are obtained with very thin paraffin sections. From water the sections are transferred to a 2-2.5 P er cent - aqueous solution of ferric ammonium sulphate for 4 to

502 HISTOLOGY

8 hours. Wash quickly in water. Stain for 12 to 24 hours in a wellripened solution consisting of 0.5 gm. haematoxylin dissolved in 10 c.c. of absolute alcohol and added to 90 c.c. of water. Wash in water. Differentiate in the iron alum solution, controlling the result under the microscope. The section should be washed in water before each examination. When the stain is limited to the nuclei, and these are sharp, wash in running water for 15 to 30 minutes; 50 per cent.; 70 per cent.; 80 per cent.; 95 per cent.; absolute alcohol; absolute and xylol; xylol.

If a counterstain is desired, add to 95 per cent, alcohol enough of a i per cent, solution of Orange G in 50 per cent, alcohol to give a deep orange color. Transfer the section from 95 per cent, to the stain for a few minutes. Wash well in 95 per cent., and pass through absolute, absolute and xylol, to xylol.

Weigert's Iron Haematoxylin.

A. Haematoxylin crystals i gm.

Alcohol, 95 per cent 100 c.c.

B. Liquor ferri sesquichlorati 4 c.c.

Water 95 c.c.

Hydrochloric acid i c.c.

At the time of using, mix equal parts of A and B. Transfer the sections from water to the stain for 2 to 5 minutes. Wash in water. If the section is overstained, add a few drops of hydrochloric acid to the water. To stop the decolorization, dip in water made alkaline with a little ammonia. This is an excellent stain and gives brilliant results. If a counterstain is desired, place sections for about i minute in Van Gieson's mixture:

Picric acid, sat. aq. sol too

Acid fuchsin, i per cent. aq. sol 10

Wash in water; 95 per cent.; absolute; absolute and xylol; xylol. For celloidin sections, 95 per cent.; oil of origanum.

Safranin.

Safranin i gm.

Absolute alcohol 10 c.c.

Anilin water 90 c.c.

Anilin water is prepared by shaking up 5 c.c. of anilin oil in 95 c.c. of distilled water and filtering through a wet filter. Dissolve the safranin in the alcohol and add to the anilin water.

Safranin is to be used after fixation in Flemming's fluid. Stain thin paraffin sections for 24 hours; wash in water; decolorize in absolute alcohol, to which has been added hydrochloric acid in the proportion 1:1000, until only the nucleus retains the stain; fresh absolute; absolute and xylol; xylol.

COCHINEAL AND CARMINE 503

STAINING IN BULK.

Embryos which are to be mounted whole or cut into serial sections are commonly stained before they are imbedded. The time given for the action of the following stains is that required in the preparation of i2-mm. pigs. Larger or smaller objects should, of course, receive a longer or shorter treatment.

Alum cochineal.

Cochineal 60 gm.

Potassium alum 60 gm.

Water 800 c.c.

Boil vigorously for 20 minutes. Cool and filter. Boil the filter paper and contents with more water for the same length of time. Cool and filter. Repeat the boiling and filtering until the powder disappears from the residue. Then put all of the filtrate together, boil for about 20 minutes and make the volume up to 800 c.c.

Stain 1 2 -mm. pigs for about 36 hours. Wash in water for 10 to 15 minutes; 50 per cent, alcohol, 20 to 30 minutes; 70 per cent., i hour; 80 per cent. Imbed in paraffin and cut in serial sections.

Counterstaining the sections in the solution of Orange G described with Heidenhain's iron hasmatoxylin will bring out the nerves beautifully. The paraffin is removed in xylol, the sections passed through absolute and xylol, absolute, and 95 per cent, to the stain. Wash in several changes of 95 per cent.; and pass back through absolute, and absolute and xylol, to xylol.

Borax carmine.

Borax 20 gm.

Carmine 30 gm.

Water 500 c.c.

Boil until everything is dissolved. Cool and add 500 c.c. of 70 per cent, alcohol. Let stand 24 hours and filter.

Stain a i2-mm. pig about 36 hours; water 10 to 15 minutes; 0.5 per cent, hydrochloric acid in 70 per cent., 30 minutes to i hour; 70 per cent, changed several times, i hour; 80 per cent, changed twice. Imbed in paraffin and cut in serial sections. The sections may be counterstained as directed under alum cochineal.

SELECTIVE STAINS. Mallory's Phosphotungstic Acid Haematoxylin.

Haematoxylin o. i gm.

Water 80.0 c.c.

10 per cent. aq. sol. of phosphotungstic acid (Merck) 20.0 c.c.

Dissolve the haematoxylin in a little water with the aid of heat; cool, and add to the rest of the solution. If the solution does not stain, it may

504 HISTOLOGY

be ripened by the addition of 10 c.c. of | per cent. aq. sol. of potassium permanganate.

Tissues must be fixed in Zenker's fluid. Sections are transferred from water to | per cent, aqueous solution of potassium permanganate for 3 to 5 minutes, washed in water, and put for 5 to 10 minutes in 5 per cent, aqueous solution of oxalic acid. Wash thoroughly in several changes of water, and stain in the haematoxylin solution for 12 to 24 hours. Transfer directly to 95 per cent, alcohol for not more than i or 2 minutes, followed by absolute for paraffin sections. Clear in xylol, using the filterpaper blotting method for celloidin sections (see p. 508).

Neuroglia, myoglia, and fibroglia fibrils and fibrin are stained blue; collagen fibrils reddish-brown; mitotic figures well shown.

Mallory's Connective Tissue Stain.

Anilin blue soluble in water (Griibler) 0.5 gm.

Orange G (Griibler) * 2.0 gm.

Phosphomolybdic acid, i per cent. aq. solution. 100.0 c.c.

Paraffin or celloidin sections of material fixed in Zenker's fluid are transferred from water to a 0.2 per cent, aqueous solution of acid fuchsin for 5 to 20 minutes. Transfer directly to the anilin blue solution and stain for 20 minutes or longer. Wash in several changes of 95 per cent, alcohol. Clear celloidin sections in oil of origanum. Paraffin sections are passed through absolute, absolute and xylol, to xylol.

Fibrils of connective and reticular tissue, amyloid, and mucus stain blue; nuclei, cytoplasm, muscle, axis cylinders, and neuroglia fibers stain red; red corpuscles and myelin, yellow.

Weigert's Resorcin-fuchsin. Boil, in an evaporating dish, 2 gm. of fuchsin and 4 gm. of resorcin in 200 c.c. of water. When it is boiling briskly, add 25 c.c. of liquor ferri sesquichlorati. Stir and boil for 5 minutes. Cool and filter. Allow the precipitate to dry; return the filter paper with precipitate to the dry dish; add 200 c.c. of 95 per cent, alcohol and boil, stirring constantly. Fish out the paper. Cool and filter; add alcohol until the volume of 200 c.c. is reached and add 4 c.c. of hydrochloric acid.

From 95 per cent, alcohol the sections, preferably fixed in alcohol or formaldehyde, are transferred to the stain for 20 minutes to an hour. Wash in 95 per cent. Clear in xylol by the blotting method (p. 508).

The elastic fibers are stained a deep purple. The remainder of the tissue should be nearly or quite colorless. If other parts are affected, the sections should be washed in alcohol containing 0.5 per cent, of hydrochloric acid. A light nuclear stain with alum haematoxylin after the elastic tissue has been stained will increase the value of the specimen.

Scharlach R. Frozen sections of fresh or formalin fixed material are stained from 15 minutes to over night in a saturated solution of the

SCHARLACH 505

dye in 70 per cent, alcohol. The sections are transferred from water to the stain, which has been freshly filtered into a tightly closing vessel. Evaporation of the alcohol causes a precipitation of the staining material. Wash in water, stain the nuclei lightly with alum haematoxylin, and mount in glycerin. Fat and lipoids stain red.

Nile Blue. Frozen sections of fresh material or material fixed in formalin for not more than 12 hours are stained for 15 minutes to 2 hours in a saturated aqueous solution of nile blue (Griibler). Wash in distilled water for 5 minutes or more, and transfer to tap water. If after 5 minutes in the tap water the section does not assume a reddish hue, add a small amount of alkali to the tap water. When the section is reddish, transfer to distilled water. Mount in glycerin or glycerin jelly and examine at once. Neutral fat red; lipoids blue.

Osmic Acid. Fat and myelin in fresh tissues may be blackened in a i per cent, aqueous solution of osmic acid. The myelin sheaths of teased nerve fibers may be so treated, the fragments dehydrated, cleared in chloroform, and mounted in chloroform damar. Sections may be prepared from tissues fixed in Marchi's fluid (p. 491), showing the fat blackened by the osmium. Use chloroform to remove paraffin from the sections, and mount in chloroform damar.

Wright's Blood Stain. After 0.5 gm. of sodium bicarbonate has been completely dissolved in 100 c.c. of distilled water, add i gm. of Griibler's methylene blue (either the form called BX, Koch's, or Ehrlich's rectified). "The mixture is next to be steamed in an ordinary steam sterilizer at 100 C. for i hour, counting the time after steam is up. The heating should not be done in a pressure sterilizer, or in a water bath, or in any other way than as stated." The mixture is then removed from the sterilizer and allowed to cool, the flask being placed in cold water if desired. When cold, it is poured into a large dish or flask. Add to each 100 c.c. of the methylene blue solution, stirring or shaking meanwhile, about 500 c.c. of a o.i per cent, solution of Griibler's yellowish eosin soluble in water. The eosin solution should be added until the mixture, losing its blue color, becomes purple, and a scum with yellowish metallic luster forms on the surface, "while on close inspection a finely granular black precipitate appears in suspension." The solution is then filtered and the precipitate allowed to become perfectly dry on the filter paper. The stain is made by dissolving 0.3 gm. of the precipitate in 100 c.c. of pure methyl alcohol. The stain need not be filtered, and like the precipitate, it keeps indefinitely. If by evaporation of the alcohol it becomes too concentrated, as shown by the formation of a precipitate when it is used, it should be filtered and a small quantity of methyl alcohol added.

Blood is obtained usually from a needle puncture in the lobule of the ear. A drop of blood is caught in the center of a perfectly clean dry

506 HISTOLOGY

cover, and another clean dry cover is immediately dropped upon it. The blood should spread between the two cover glasses, forming a film which cannot be too thin. The covers are then drawn rapidly apart (they should slide along one another and not be lifted apart). The blood film dries from exposure to the air, and remains stainable for weeks.

To stain the blood film, the cover glass is to be held in cover-glass forceps with the film side uppermost. The stain is applied as follows:

1. Cover the preparation with a noted quantity of the stain by means of a drop-bottle or medicine dropper.

2. After i minute add to the staining fluid on the preparation the same quantity of distilled water, by means of the dropper, and allow the mixture to remain i\ minutes, not longer. Longer staining may produce a precipitate. The total quantity of diluted fluid on the preparation should not be so much that some runs off. A metallic scum forms when the stain is properly diluted, but the stain should not become transparent.

3. Wash the preparation in tap water for 30 seconds, or until the thinner portions of the film become yellow or pink. Disregard the thick parts, which are blue. The process of decolorizing may be watched through the microscope by placing the cover glass with film side uppermost on a slide.

4. Dry and mount directly in damar.

Silver Nitrate. Intercellular cement spaces and the boundaries of endothelial cells may be blackened by a i to i per cent, aqueous solution of silver nitrate, which acts chiefly upon free surfaces. The fresh tissue should be kept flat, the mesentery, for example, being tied over a detached bottle neck, while it is immersed in the solution for i to 10 minutes. Transfer to distilled water, and expose to direct sunlight. As soon as it becomes brown, usually in 5 to 10 minutes, it is washed in 0.6 per cent, salt solution. If desired, the nuclei may be lightly stained in alum haematoxylin. Examine in glycerin, or dehydrate clear in xyol and mount in damar.

Blood vessels may be injected through glass tubes with the silver solution. Sections are made and exposed to the light, and the outlines of the endothelial cells become dark.

5. Clearing and Mounting. CLEARING.

Before satisfactory permanent preparations can be obtained, the sections or object must be cleared,. This is accomplished by infiltrating the tissues with substances which, by reason of their high index of refraction, render the tissues more or less transparent. Structures to be studied are previously stained and thus easily rendered prominent.

CLEARING AGENTS 507

A variety of reagents with widely different chemical properties are used. Glycerin and acetate of potash are commonly used for frozen sections or teased preparations which, for any reason, cannot be mounted in damar.

The choice of a clearing agent for damar mounts depends chiefly on two factors, the kind of stain employed and the imbedding medium.

Xylol. This is the best clearing agent for use after aniline dyes. It clears only from absolute alcohol, through which celloidin sections cannot be passed, since it dissolves celloidin. However, it can be used for celloidin or other sections dehydrated in 95 per cent, alcohol by the following method. Blot the section on the slide with smooth, soft filter paper and pour on a few drops of xylol. Repeat the blotting, followed by xylol two or three times and the section will be perfectly clear. Paraffin sections attached to the slides are cleared by immersion in a vial of xylol; this has already been mentioned as the last step in the staining processes.

Oleum Origani Cretici. This clears readily from 95 per cent, alcohol without dissolving celloidin and affects aniline colors slowly. Although particularly recommended for clearing after celloidin imbedding, it is useful for all kinds of sections.

Carbol-xylol.

Carbolic acid crystals i

Xylol 3

Used for clearing thick sections of the central nervous system after carmine and haematoxylin stains. Clears from 95 per cent, alcohol without affecting celloidin, but extracts the basic aniline dyes.

Chloroform. Since osmic acid reduced by fat is soluble in xylol, chloroform is used in cases where permanent mounts of such preparations are desired (as after fixation in Marchi's fluid).

MOUNTING.

Frozen sections which cannot be mounted in damar are mounted in glycerin, potassium acetate or glycerin jelly.

Glycerin Jelly. Soak 7 gm. of gelatin for about 2 hours in 42 c.c. of distilled water. Add 50 gm. of glycerin. Warm, stirring constantly for icr-i5 minutes. Filter hot through moistened cotton.

Bring sections on a slide and blot off excess water. Put on a small piece of glycerin jelly and warm gently until it melts. Cover and cool.

Preparations mounted in any of these three substances may be rendered more or less permanent by coating the edges of the cover and adjacent surfaces of the slide with paraffin or wax.

Gum Damar. Of the two substances most commonly employed for permanent mounts, namely, Canada balsam and damar, the latter is preferable, since balsam turns yellow with age.

508 HISTOLOGY

Colorless pieces of damar are dissolved in xylol and filtered. If the solution is too thin, evaporate to the proper consistence; if too thick, add more xylol. The proper consistence is that of a thin syrup.

A solution in chloroform should be prepared in similar manner for use with osmium preparations.

After paraffin sections have been cleared (3 to 5 minutes), the excess of clearing agent is drained away and the surface of the slide outside the sections is wiped off. The section must not be allowed to become dry. A drop of damar solution is placed at once upon the section and a cover glass is carefully lowered over it. With all preparations, whatever the mounting medium may be, the cover glass should be lowered in the following manner. It is held over the specimen and its left edge is first brought into contact with the slide; a needle held in the left hand holds this edge in position. Another needle held in the right hand with its point beneath the right edge of the cover enables one to have perfect control of the cover glass while it is being lowered. The contact between the cover glass and the mounting medium spreads gradually from left to right as the cover is lowered, expelling the air as it advances. If air bubbles are caught in the medium, the cover may be alternately raised and lowered a little until they escape, but once the cover is flat upon the specimen it should not be lifted. The cover glass should be somewhat larger than the specimen so as to extend beyond it on all sides.

Celloidin sections which have been cleared in oil are floated over the blade of a spatula placed in the oil, and are spread out flat upon it with the aid of a needle. They are then transferred to a clean glass slide, being pulled from the spatula with the needle. They should be moved to the exact center of the slide, if the preparations are to look well, and then the oil is removed by placing two thicknesses of filter paper over the section and pressing upon it quite firmly; at the same time the section is made smooth. A drop of damar is then placed upon the section before it dries, and the cover glass is applied as described in the preceding paragraph.

Damar mounts are then labelled and may be placed in a drying oven with a temperature of 35-40 C. They may be used after a few days, but the damar is not thoroughly hardened for a considerable time.

SLIDES AND COVER GLASSES.

Slides should be of colorless glass with ground edges. For ordinary use, slides measuring i X3 inches (26 X 76 mm.) are preferable. Occasionally, as in mounting serial sections, or large sections of the central nervous system, wider slides are needed. Thick slides are preferable to thin ones.

COVER GLASSES 509

For ordinary use, cover glasses 18-22 mm. square are sufficient. Occasionally, as in mounting serial sections or large specimens, oblong covers may be needed. If possible, no covers ranging outside of 0.150.18 mm. in thickness (No. i grade) should be used, since thicker covers (No. 2 grade) often prevent the oil immersion lens from being brought into focus. Many valuable sections have been destroyed in attempting to focus through thick cover glasses.

Clean the slides and covers by dipping in alcohol and drying with a soft crash towel or old linen handkerchief. Sometimes it may be necessary to wash them in 10 per cent, nitric acid, followed by a thorough washing in water and then in alcohol. Slides which remain hazy after thorough washing must be discarded.

INJECTIONS.

The courses of blood and lymphatic vessels and of ducts are studied by means of injections. Transparent, deeply colored fluid mixtures, which will harden in the vessels, are used. So-called "warm" injection masses, which contain gelatin, give more perfect results but are more difficult to use than "cold" injection masses.

A tapering glass or metal cannula is inserted into the vessel or duct, which is then tied securely around it. From a syringe connected with the cannula by a short rubber tube, the mass is then forced into the vessel. Pressure may also be obtained by having the injection mass in a receptacle which is connected with the cannula by a long flexible tube. The pressure is varied to suit the needs of the moment by raising and lowering the receptacle.

When a warm injection mass is being used, the bottle containing the mass must be placed in a water-bath and kept at a temperature of about 45 C. The organ or animal to be injected must also be placed in a water- bath of the same temperature.

Organs to be injected must be perfectly fresh; they may be left within the body or removed and injected separately. It is advisable to wash out blood vessels with warm salt solution or Ringer's solution before the injection.

It is important that in connecting the end of the tube carrying the injection mass with the cannula inserted into the vessel, no air bubbles be allowed to enter.

COLD INJECTION MASSES. i. Blue Injection Mass.

Soluble Berlin blue i

Distilled water. . . .20

510 HISTOLOGY

2. Carmine Injection Mass. Dissolve i gm. of carmine in i c.c. of strong ammonia plus a little water; dilute with 20 c.c. of glycerin. Add i gm. of common salt dissolved in 30 c.c. of glycerin. To the whole solution add an equal quantity of water.

WARM INJECTION MASSES.

1. Berlin Blue. Allow clear sheets of best French gelatin to swell up for i or 2 hours in double the quantity of water. Dissolve by warming gently over a water-bath and add, stirring constantly, an equal volume of a warm solution of Berlin blue prepared as directed above. Filter through flannel wrung out in hot water.

2. Carmine. This is the best injection mass to use, but it is very difficult to prepare. Dissolve 2 to 4 gm. of the best carmine in the required amount of ammonia. Filter and stir into 10 to 50 gm. of a filtered warm solution of gelatin, prepared over the water-bath as described above. Then add 25 per cent, acetic acid, drop by drop, stirring .constantly, until the mass becomes bright red and loses its ammoniacal odor. If too much acetic acid is added a precipitate forms and the mass is spoiled. Filter through warm flannel.

Organs injected with a cold mass are put into 80 alcohol. After a few hours they may be cut into pieces. After injection with a warm mass, the specimen is put into cold water to hasten the solidification of the gelatin, and then transferred to 80 per cent, alcohol. Imbed in celloidin. Thick sections are necessary in order to follow the course of the vessels.

Prepared injection masses for use, cold or warm, are sold by dealers in microscopical supplies.

Many ingenious injection methods have been devised, such as the injection of small living embryos by allowing ink to enter the veins and be distributed through the body by the action of the heart; and vessels have been injected with milk, after which frozen sections were stained with Scharlach R.

SPECIAL METHODS.

The following special methods are included because of their fundamental importance. For the many other special methods which are occasionally of service, reference should be made to the works on technique mentioned at the beginning of this section.

Weigert's Method for Staining Myelin Sheaths. This is a method for the differential staining of the myelin sheath of nerve fibers and is much used in the study of the normal and pathological histology of the central

WEIGERT'S MYELIN STAIN 511

nervous system. As a result of some chemical reaction between myelin and a chrome salt, the myelin is fixed so that it does not dissolve in alcohol and ether, and at the same time is mordanted so that it stains deeply with haematoxylin.

1. Fix in a 10 per cent, aqueous solution of formalin for several days to several weeks, or indefinitely. Use a large quantity of fluid; change at the end of 24 hours and thereafter whenever it becomes cloudy.

2. Cut the tissue into pieces not over i cm. thick, and place in a 2.5 per cent, aqueous solution of potassium bichromate renewed each day for 3 or 4 days; then in a 5 per cent, solution renewed each day for 3 to 4 days.

3. Wash in running water for 24 hours (large pieces several days).

4. Transfer the tissues to the following solution (Weigert's second mordant) for 24 to 48 hours.

Acetate of copper 5.0 gm.

Acetic acid, 36 per cent, solution. . 50 c.c.

Fluorchrom 2.5 gm.

Water 100. o c.c.

Boil the fluorchrom and water in a covered dish; turn off the gas and add the acetic acid and then the acetate of copper. Stir until the latter dissolves, and cool.

5. Wash in running water 24 hours or longer.

6. Dehydrate in graded alcohols and imbed in celloidin. Cut sections 20-25 p thick.

7. Stain sections for 12 to 24 hours in:

Ripened 10 per cent, solution of hsmatoxylin in absolute alcohol 10

Saturated aqueous solution of lithium carbonate i

Water ; 90

The haematoxylin is kept as a stock solution and combined with the carbonate of lithium and water at the time of using. Beautiful results may also be obtained by staining over-night or longer in Weigert's ironhaematoxylin (p. 502).

8. Wash in water.

9. Differentiate in the following solution:

Borax 2.0

Potassium ferricyanide 2.5

Water 100.0

It is advisable to dilute this solution with i or 2 volumes of water. After the first staining method, the gray substance of the sections appears yellow; after the iron-haematoxylin it is colorless.

10. Wash in running water 4 hours to over-night.

u. 95 per cent, alcohol, 3-5 hours (may be left over-night)

12. Fresh 95 per cent., 5 minutes.

13. Clear in carbol-xylol.

14. Mount in xylol damar.

512 HISTOLOGY

Pal's Modification of Weigert's Stain. The tissue is fixed, mordanted and imbedded as directed in i, 2, 3 and 6 above. Sections may be very much thicker.

7. Sections are placed for several hours in a per cent, aqueous solution of chromic acid, or for a longer time in a 2.5 per cent, solution of potassium bichromate. (May be omitted.)

8. Stain for 24 to 48 hours in:

Ripened 10 per cent, solution of haematoxylin in absolute alcohol 10

Water 90

9. Wash in water plus i to 3 per cent, of a saturated aqueous solution of lithium carbonate until the sections appear of a uniform deep blue color.

10. Differentiate for 20 seconds to i minute in a \ per cent, aqueous solution of potassium permanganate.

11. Place for a few seconds in the following solution, until the gray substance is colorless or nearly so:

Oxalic acid i

Potassium sulphite i

Water 200

12. Wash in water.

Repeat steps 10, n and 12 until the differentiation is complete. Then wash 4 hours or longer in running water.

13. 95 per cent, alcohol, 3 to 5 hours.

14. Fresh 95 per cent., 5 minutes.

15. Carbol-xylol. Mount in xylol damar.

Golgi's Method for the Impregnation of Nerve Cells. This method depends on the formation of a fine precipitate in certain tissue elements or in pre-existing spaces when the tissues are treated with a solution of potassium bichromate and then with a solution of silver nitrate or mercuric bichloride. The value of the method lies on the fact that it picks out here and there a cell and stains it with its processes more or less completely. This same fact renders the method very uncertain. Of the several modifications of this stain, only one the so-called rapid method is here given.

1. Pieces of fresh tissue about 5 mm. thick are placed for 3 to 8 days in the following solution:

Osmic acid, i per cent, solution i part

Bichromate of potassium, 3 . 5 per cent, solution 4 parts

2. Transfer to a large quantity of 0.75 per cent, solution of silver nitrate for 2 or 3 days.

Keep the tissues in the dark during treatment with both fluids. The length of time the tissues should remain in the first solution depends on the elements it is desired to impregnate.

WEIGERT'S MYELIN STAIN 513

For the human cord, the time is approximately as follows: for neuroglia, 2-3 days; nerve cells, 3-5 days; for nerve fibers and collaterals, 5-7 days.

3. Cut sections 50 to 100 p thick. They may be made free-hand or with a microtome. Blocks may be quickly imbedded by dehydrating for a few minutes in absolute alcohol, and placing in a thick solution of celloidin for about 5 minutes. Harden in 80 per cent.

4. Sections are dehydrated quickly in alcohol.

5. Clear in oil of cloves or bergamot.

6. Mount without a cover glass in xylol damar and dry quickly at 40 C. Protect the sections from the light and dust as much as possible.

If the method is unsuccessful, the specimens may be transferred back to an osmic acid and bichromate mixture containing less osmic acid, and after several days again placed in the silver nitrate solution for 24 to 48 hours.

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Pages where the terms "Historic" (textbooks, papers, people, recommendations) appear on this site, and sections within pages where this disclaimer appears, indicate that the content and scientific understanding are specific to the time of publication. This means that while some scientific descriptions are still accurate, the terminology and interpretation of the developmental mechanisms reflect the understanding at the time of original publication and those of the preceding periods, these terms, interpretations and recommendations may not reflect our current scientific understanding.     (More? Embryology History | Historic Embryology Papers)

Histology with Embryological Basis (1913):   Part I. 1.1. Cytology | 1.2. General Histology | 1.3. Special Histology Part II. 2.1. The Preparation of Microscopical Specimens | 2.2. The Examination of Microscopical Specimens

Reference: Lewis FT. and Stöhr P. A Text-book of Histology Arranged upon an Embryological Basis. (1913) P. Blakiston’s Son and Co., 539 pp., 495 figs.

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Cite this page: Hill, M.A. (2024, April 25) Embryology Book - A text-book of histology arranged upon an embryological basis (1913) 2-1. Retrieved from https://embryology.med.unsw.edu.au/embryology/index.php/Book_-_A_text-book_of_histology_arranged_upon_an_embryological_basis_(1913)_2-1

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