PART II.

THE TREATMENT OF THE MATERIAL.

INTRODUCTION. The material obtained by autopsy, surgical operation, curettage, excision, spontaneous discharge, animal experimentation, etc., may be examined microscopically in the fresh state, or prepared for microscopic examination by methods of fixation, hardening and imbedding. The methods necessary for such histologic studies of pathologic material are given in the following pages, arranged as far as possible in their logical order. Only those methods are given that, in the light of the writer’s experience, yield the best results, from the standpoints of economy of time, labor, and expense, and perfection of result. The number of histologic methods contained in the literature is so great that it is out of the question for the student or practical worker to try out all of them. To give all of these methods would create confusion. I have attempted to avoid this by giving in full detail only those which in our laboratory experience have yielded the best results. So many methods represent but slight variations of some original method, and in the great majority of cases these variations add so little or nothing of value to the original method that in such cases the latter alone is given in full, with references only to the variations of the method. The individual equation plays such a large part in the judgment of laboratory methods that allowance has been made for this when certain variations or alterations in original methods have been strongly recommended by expert laboratory workers.

The purpose of the microscopic examination is the revealment of pathologic changes too small to be recognized by the naked eye, and the securing of a diagnosis that cannot be made macroscopically, as well as the confirmation of diagnoses based upon the gross appearances. Aside from these more immediate practical considerations, the microscopic examination of tissues is concerned with the solution of etiologic and pathologic problems, and the extension of our knowledge of disease. The aim of pathologic technique is the fixation of tissues for microscopic examination in such a manner that all of the morphologic and chemical elements and constituents of the tissue are perfectly preserved, so that with differential staining methods they are all brought out with sufficient contrast to be readily and correctly identified. In a certain number of methods this ideal is attained, and to Weigert, more than to any other worker in the field of pathologic technique, are we indebted for such ideal methods.

The choice of methods will depend upon the source and nature of the material, the object of the examination, the time-element and the degree of responsibility involved. The cellular elements of all pathologic fluids, secretions and excretions should be examined in the fresh state as well as in fixed preparations. For the demonstration of various chemical and morphologic features that are lost or altered by processes of fixation and imbedding, and when a rapid diagnosis is required, the examination of the material in its fresh state or by the freezing method is indicated. When the freezing method cannot be employed because of the changes in cells and tissue produced by it, when very thin or serial sections are desired, when a rapid diagnosis is not required, and when a very careful and minute study is desired, with the application of various staining methods, then the material should be fixed, hardened and imbedded and cut upon a microtome. Whatever method is chosen, it must be borne in mind, particularly in practical diagnostic work, that the portions chosen for microscopic examination must represent the characteristic anatomic structures of the tissue or organ, that living tissue be included, that the pathologic condition be represented both in its fully-developed state and at the transition-border between it and the healthy tissue, and that when sections are cut the block or tissue must be so oriented as to give the most comprehensive view of the tissue and the pathologic process. To accomplish this fully it is often necessary to make a number of blocks representing different areas of the material, and to cut these in different planes.

CHAPTER XVII.
THE LABORATORY OUTFIT.

For practical diagnostic work or for pathologic research various instruments and utensils are necessary, although the expense of fitting up a working pathologic laboratory is not as great as it is often thought to be. The most expensive item, as well as the most important, is the microscope. This should be of the best make, and should be carefully selected and tested before the final purchase. As a rule the German makes, Zeiss, Leitz, and others, are to be preferred to the American instruments, in spite of the higher cost due to the duty imposed. I have found the German microscopes uniformly good and standing the wear and tear of a teaching laboratory much better than the American-made stands. I have never seen a poor Zeiss or Leitz objective, but cannot say the same thing of other makes. On the other hand, I have seen some American objectives that were as good as any German ones, but there are not many such. If one is going to buy an American microscope it should be bought on the same principle that one would buy a violin or a piano, wholly on its individual merits; and these can be ascertained only by having the instrument carefully examined and tested by an expert. Most laboratory workers will agree that the Zeiss instruments are the best; they are also the most expensive. For all practical purposes a Leitz stand costing ninety to one hundred dollars is quite good enough. A medium-sized continental stand, with rack and pinion and micrometer screw for coarse and fine adjustment, a triple nose-piece with dust-protector, Abbé condenser, iris diaphragm, plane and concave mirrors, three objectives (low, high and 1/12 oil-immersion), and two eye-pieces, a low and a high, form a complete outfit that answers all practical requirements. The new type of stand with curved arm and large stage, permitting the examination of all parts of a Petri dish or glass plate, and with the mechanism of the fine adjustment protected from any strain when the instrument is lifted by the arm is especially recommended. I have also found the black-finish very practical. The entire outfit need not be purchased at once; the stand with its accessories and a low power may first be purchased, and the higher-power objectives obtained later. One of the first luxuries is a movable adjustable stage. A very good and relatively cheap one is made by the Spencer Lens Co., of Buffalo. If a Zeiss stand is purchased the objectives A, D, and 1/12 oil-immersion, and oculars 2 and 4 best meet the requirements. Of the Zeiss apochromatic series, the objectives 16.0, 8.0, 4.0, and oil-immersion 2.0 mm., apert. 1.30, and oculars 4, 6 and 8 are most serviceable. The apochromatic objectives and the compensation-oculars are expensive, and need not be used for ordinary work, but are indispensable for photographic work. The Leitz objectives, 3, 6, or 7, and 1/12 oil-immersion, with oculars 2 and 4, and the equivalent objectives of the Spencer Lens Co. or Bausch and Lomb will answer all ordinary needs. For the purpose of microscopic measurements an ocular micrometer is necessary. This may be obtained as a separate eye-piece, or as a round piece of glass with measured divisions marked upon it that may be put into an ocular. The value of the scale must be determined for every lens and tube-length by estimating the number of its parts covering one part of a stage-micrometer marked in hundredths of a millimetre.

Fig. 50.—A satisfactory outfit for the working laboratory. Continental stand, medium-sized, with large stage, three objectives, etc.

An instrument of any one of the above-mentioned makes, carefully selected and tested, should last its owner a life-time if proper care is taken of it. It should receive the same careful attention accorded a good violin or piano. It should be protected from dust, action of chemicals, heat, sunlight, and rough usage. When carried it should be supported in such a way that its weight is not thrown upon the thread of the adjusting screws. The adjustment, draw-tube and iris diaphragm should be carefully oiled at intervals, using the least possible amount of the best microtome oil. It is not necessary here to enter into the construction and theory of the microscope, as this knowledge has usually been obtained before the pathologic laboratory is reached. Experience has shown me, however, that it is always necessary to remind students, even those experienced for some time in the use of the microscope, of certain fundamental principles in the adaptation of microscopic technique to pathologic work. The following rules are of value:—

1. Use a low-power objective for all work except for the study of bacteria, microparasites and finer cell-structures. The aim should be to obtain as much of a bird’s-eye view of the “geography” of the section as possible. Contrast plays a very important part in pathologic diagnosis; and it is lost in high-power work, so far as the relations of cells and tissue-elements and pathologic products are concerned. The student almost invariably enters the pathologic laboratory with a fixed “high-power habit,” and he is usually greatly surprised to learn how much he misses with the high-power and how much he can see with the low-power. A motto used many years in my laboratory, “Low-power objective and high-power cortex,” is of greater educational value than may appear at first sight. A slide should be examined first with the naked-eye, as it is held against a window or light; then it should be examined under the low power. Rarely will it be necessary to use a high-power except for the purposes mentioned above.

2. Weak eye-pieces should be used; strong ones darken the field and tire the eyes. Only in the case of apochromatic lenses and compensation oculars can the strong ones be used without darkening the field too much.

3. In the use of higher powers see that the tube is drawn out to the proper length, as indicated in the directions sent with the instrument.

4. For pathologic work an Abbé condenser is essential. It should be pushed up to its proper position beneath the stage, and the plane mirror should be used with it, reflecting the light from a cloud if possible. Daylight is always the best light. When this cannot be obtained an incandescent or Welsbach lamp with ground-glass globe can be used. The concave mirror should then be used. The yellowish tint of artificial light may be avoided by the use of a piece of blue glass placed beneath the condenser, or a vessel of copper sulphate solution may be interposed. Especial lamps designed to meet the requirements are offered by the trade.

5. The iris-diaphragm should be adjusted by the same hand that moves the slide, usually the left one. With unstained preparations the diaphragm should be nearly closed; when using the oil-immersion it should be fully opened, as is also the case when stained preparations are studied with the lower power. With higher magnification the aperture is diminished somewhat, although color-effects are best shown with open diaphragm. In the study of pigments the diaphragm should be fully closed for a few moments to see if the pigment shows any color by reflected light. It is then examined by open diaphragm. In the study of sections the mirror and diaphragm should be manipulated in various ways to bring out all of the detail of the preparation, and should be adjusted to suit each preparation.

6. Objectives must never be screwed down until they strike the slide or stage. The higher-powers are frequently ruined in this way. When running the objective down always examine from the side to see that there is no danger of its striking the stage. In the use of the oil-immersion place the drop of oil upon the slide or cover-glass, and lower the objective by turning the coarse adjustment until the oil spreads out between the lens and the glass; then focus with the fine adjustment until a well-defined field is obtained. The oil-immersion lens should not be allowed to stand many consecutive hours in the oil. The oil should be cleaned from the lens by wiping the latter with lens-paper or a soft cloth; if the lens is sticky the paper or cloth may be moistened with benzol. The lens itself should never be wet with benzol, xylol, alcohol or any cleaning-fluid, because of the danger of softening the balsam in which the lenses are imbedded.

7. Use the mechanical stage only for differential blood-counting, or when the entire section is to be gone over carefully, or when certain details are found with difficulty and it is desirable to mark them for future reference. An immense amount of time is lost in the use of the mechanical stage for ordinary work. By moving the slide with the fingers of the left hand resting upon the stage an entire section may be gone over in a few seconds without missing any part of it; to accomplish the same thing with the mechanical stage requires much more time.

It is an excellent plan for the student to purchase his microscope when entering the medical school and to use his own instrument throughout his course. It is the one instrument without which no physician can afford to enter practice; and the student who uses his own microscope before graduation will continue to use it afterward. The microscope obtained, the remaining expenditure necessary for the fitting-up of a practical working laboratory of clinical and pathologic diagnosis need not be very great if one’s financial condition does not warrant spending with a free hand. It is possible with a little labor and ingenuity to make at home, or to show the local tinsmith how to make, a large part of the necessary apparatus, such as sterilizers, paraffin-ovens, drying ovens, thermo-regulators, etc., at a slight cost. Students of mine have made these things out of old tin cans and glass tubing; one student at a cost of less than three dollars constructed a microtome on which practical working sections could be cut. For the celloidin method no apparatus except the microtome is necessary, as the process of imbedding is carried on in bottles or dishes. These points are mentioned to offset the prevalent idea that a large expenditure is a necessity in installing a practical working laboratory.

In a large diagnostic laboratory, or in one intended for teaching and investigation, there are numerous accessories necessary to modern microscopic technique. For the observation of living objects a warm stage is needed. The simple electrical apparatus devised by Ross is the most convenient form, as it can be slipped on and off the slide without changing the focus. It can be attached to any electric light circuit and requires no attention.

For drawing from the microscope the improved form of the camera lucida, or the latest model of the Edinger drawing-apparatus are recommended. Both of these instruments have recently been greatly improved. The Zeiss microphotographic apparatus is by far the best for microphotographic work. For the polarization-microscope, microspectroscope, and the complicated and expensive ultra-violet and dark-field apparatus the worker is referred to the Zeiss catalogues. A simple and practical dark-field method for the illumination of bacteria, spirochætes and ultramicroscopic particles suspended in fluids requires only a strong illumination and the use of a Zeiss, Leitz, or Reichert dark-field condenser; or the very simple “India-ink” method may be used for the demonstration of spirochætes. (See Staining of Spirochætes.) Especial instruments for easily finding a certain field are obtainable, and are of great convenience in marking slides for photographic purposes.

Fig. 51.—A good practical microtome for paraffin and celloidin work. Well-adapted to needs of students and beginners.

Next to the microscope the most important instrument in pathologic work is the microtome. One that can be used for either celloidin or paraffin work, and that can also be utilized as a freezing microtome, should be selected in private work, when economy is desired. The majority can be used for either paraffin or celloidin, and one of the Becker models can be easily attached to the carbonic acid holder for the cutting of frozen material. The Bardeen freezing microtome is relatively inexpensive, very satisfactory, and can be easily attached to the carbonic acid tanks. It can be recommended for freezing work. The “slide” type, either that in which the knife-holder is moved by the hand directly or by a crank turned by the hand, is advised for ordinary diagnostic work. For students and beginners the crank is of great advantage, as the knife is securely held and cannot jump. The best microtomes are made by Schanze of Leipzig, Jung of Heidelberg, Becker of Göttingen, the Cambridge Scientific Co., the International Instrument Co., and the Bausch & Lomb Co. For either celloidin or paraffin work the medium or large Schanze slide-microtome or the Minot’s precision microtome are recommended; for cutting serial sections in paraffin the latest modification of the Minot automatic rotary microtome is especially adapted. The best microtome knives are made by Walb of Heidelberg. A long, heavy knife is to be preferred to a light one. For the freezing microtome a knife of the type of the blade of a carpenter’s plane set in a wooden handle should be used, Hones and strops of the best quality are necessary.

Paraffin-ovens and drying-ovens of suitable size, and constructed preferably of copper, are necessary for paraffin work. The water-space about the oven should be sufficiently large, and the temperature should be controlled by a thermo-regulator. Various models are offered in the trade, but they can be made more cheaply by the local tinsmith. The thermo-regulator can also be home-made by anyone who has the necessary training in glass-blowing usually given in courses in bacteriology.

Various instruments and utensils, such as razors, double-bladed knives, forceps, spatulas, section-lifters, needles, scalpels, scissors, glass rods and tubing, test-tubes, graduates, flasks, funnels, bottles, staining dishes, reagent bottles, rubber tubing, water-bath, tripods, centrifuge, Bunsen burners, asbestos pads, gauze, filter-papers, absorbent paper, labels, oil and wax colored pencils, slides, cover-glasses, slide-boxes, camel’s-hair brushes, glass droppers, platinum wire, etc., are required for the pathologic laboratory, and can be chosen to suit the individual needs. The solid watch glasses make very good small staining dishes; the enamelled trays and glass dishes used in photographic work are especially adapted to the plate-method, particularly the size used for the 4 × 5 plate. Tea-strainers or small sieves can be used for staining a large number of celloidin sections; and there are different types of staining-dishes designed for the staining of slide- and cover-glass preparations in number. Slides should be of the best quality, colorless and with ground edges, and of medium thickness. Cover-glasses should be square or oblong, round covers having but little use in pathology. For ordinary work the No. 2 square, ¾ inch, is recommended; for work with the higher-power dry objectives a thinner cover must be used. Slides and covers may be cleaned by placing them in a solution of equal parts of one per cent sulphuric and chromic acids and then rinsing in distilled water. A good cleaning fluid is also made of one part acetic acid to three of 80 per cent alcohol. To clean old mounts melt the balsam by heat or dissolve in turpentine, separate slides and covers, boil in ten per cent lysol for half an hour, or for ten minutes in the sulphuric-chromic-acid mixture, rinse thoroughly, dry with cloth having no lint.

The laboratory should be supplied with running water, a sink large enough for washing out specimens, numerous stop-cocks, and a drip-board. Distilled water in abundance must be available. The laboratory-table should have an alcohol- and xylol-proof finish, black on the whole being the most practical color. A portion of the table should be covered with glass beneath which there is laid a sheet of white paper.

CHAPTER XVIII.
THE EXAMINATION OF FRESH MATERIAL.

I. METHODS OF EXAMINATION.

Pathologic material is examined in the fresh state when it is desired to make a diagnosis in the shortest time possible, or when the processes of fixation and hardening produce such alterations in the morphology and chemic constitution of the cells that these features can be recognized only in the unfixed, fresh state. So far as the saving of time is concerned it is possible to take material removed during an operation, examine it in the fresh state, and return a diagnosis to the surgeon, while the patient is still on the table under the influence of the anæsthetic. It is not possible to do this with all tissues or with all pathologic conditions; but, when it can be done, the advantages of such a rapid diagnosis, in the saving of time, labor, expense and danger to the patient, are obvious. The best idea of the cell is also gained by its study in a fresh condition. Vital phenomena and certain morphologic features, as cilia, can be observed only in fresh material. Many of the chemic constituents of cells (glycogen, fat, mucin, pseudomucin, albumin-granules, cholesterin, etc.) are either lost, or are so changed by processes of fixation or hardening that they can no longer be recognized. The majority of specific chemic tests can be made in fresh tissues only. Moderate and slight degrees of fatty degeneration and cloudy swelling are easily recognized in the fresh state; in fixed and hardened preparations they may not be recognized at all. Particularly in the case of the heart-muscle is it necessary to make an examination of the fresh material when the diagnosis of these conditions is concerned. Further, a greater or less degree of shrinking is caused by many of the agents used in fixing and hardening, and this is avoided by the examination of the fresh material. In the case of pathologic fluids (sputum, urine, féces, etc.) an examination of the sediment in the fresh state should always be made as a matter of routine. The formed elements of these fluids are best determined by this means. In the case of tissues, a diagnosis made by means of scrapings, smears, teased bits of tissue, frozen sections, etc., should always be controlled by the examination of fixed and hardened material.

In the examination of fresh material the following methods are employed: Sedimentation, smears, scraping, crushing, teasing, maceration, sections, shaking or penciling, digestion, intravital staining, injection, the warm stage and “cultivation.”

1. Sedimentation. The formed elements of pathologic fluids (urine, sputum, pus, blood, exudates, transudates, cyst-contents, etc.) are examined by collecting the sediment of such fluids from the bottom of a sedimenting glass or bottle, by means of a capillary pipette controlled by the finger. While the sediment is passing up into the tube the pipette should be moved about the bottom of the vessel so as to get some of the sediment from all parts. When the fluid is rich in cellular elements sedimentation is not necessary; a drop of the fluid is placed upon the slide; if too thick it is diluted with physiologic salt-solution or serum. If poor in cellular elements the fluid must be centrifugalized by means of a water- or electric-centrifuge; and a drop of the sediment in the centrifuge tube is then removed by the pipette and placed upon the slide, and covered with a cover-glass. To facilitate the low-power examination of such sediments parallel streaks upon the slide may be made across its entire length, and examined without the use of cover-glasses. To apply the various reagents mentioned below it becomes necessary to use a cover-glass as directed.

2. Smears. A clean fresh cut is made into the organ or tissue, and a clean slide or cover-glass is drawn across the surface. Without permitting the smear to dry a drop of salt-solution or any desired reagent is put upon it, and it is then examined. This method is especially applicable to the study of the cells of the spleen, bone-marrow, lymphnodes, etc. Permanent balsam-mounts may be made of such smears by fixing with heat or alcohol and ether, staining, drying and mounting.

3. Scraping. A fresh cut is made into the organ or tissue, and the excess of blood absorbed by a pad of absorbent paper devoid of lint. A clean scalpel held at an angle of 45° is then drawn with some force back and forth over the cut-surface until its blade collects a sufficient amount of “tissue-juice” made up of the cells of the tissue. This is then put upon a slide, and covered with salt-solution or any desired reagent, and is then examined. This method is used especially for cellular infiltrations, soft tumors, and parenchymatous organs (spleen, lymphnodes, bone-marrow, liver, etc.), and for the inner wall of cysts (echinococcus-cysts, cysts lined with ciliated epithelium).

4. Crushing. A small bit of the tissue is cut out with the scissors or scalpel, and placed upon the slide. A cover-glass is then placed upon it and pressed down so firmly that the bit of tissue is spread out in a thin film or layer beneath the cover-glass. Reagents are introduced beneath the cover-glass, as desired. (See below.) This method is used in the examination of the lung, kidneys and brain for fatty embolism, and of the brain and spinal-cord for “fat-granule” cells, pigment, calcified ganglion-cells, Negri bodies, etc. It is also frequently used in the bacteriologic examination of tissues (crushing of tubercles, etc.).

5. Teasing. A small bit of the tissue is cut out and placed upon the slide or in a staining-dish and covered with physiologic salt-solution. It is then teased with fine needles until divided into its ultimate elements. Hard tumors (mature connective-tissue tumors, fibrosarcomata, etc.), muscle (examination for trichina), nerve-trunks, etc., are best examined in this way.

6. Maceration. In the case of some tissues the ultimate histologic elements are so firmly held together that they cannot be separated without the aid of a macerating- or dissociation-fluid. (See below.) Digestion is also used for the same end. The tissue should be as fresh as possible and cut into small bits, which are placed in the maceration-fluid in watch-glasses or staining-dishes for twenty-four hours or longer. The macerated bits are then teased until the finest elements are separated. In the case of very minute elements the teasing may be carried on under a hand-lens or the stereoscopic binocular microscope. During the process of maceration all parts of the macerating tissue must be kept covered with the macerating-fluid, or the uncovered portions will become hardened.

7. Section-cutting. Sections of fresh tissue may be made with the curved shears, simple razor, double-bladed razor and the freezing-microtome.

a. Curved Shears. The tissue is put upon a stretch, and from the surface a thin, flat section is cut out with the scissors. With care a fairly thin section may be obtained in this way. It may be examined by pressing it upon a slide beneath a cover-glass, or it may be treated as a frozen section.

b. Simple Razor. As a part of their required laboratory training medical students should learn how to make working sections with a simple razor. Such a technical knowledge is sure to be of practical use at some time or other. With a little practice sections sufficiently thin for ordinary diagnostic work can be cut. If the piece of tissue is large it may be held in the hand; but when small or soft it may be placed in a matrix of hardened liver, pith, potato, apple, firm lard or butter, paraffin, etc., and cut at the same time with the latter. Both blade and tissue should be wet with physiologic salt-solution. The blade, which must be very sharp, should be drawn through the tissue by a shoulder movement, with the wrist-joint fixed. As the sections are cut they are floated off of the razor-blade into physiologic salt-solution, and thence treated as desired.

c. Double-bladed Knife. This consists of two parallel blades arranged so that the space between them can be changed by means of screws. For firm material the blades should be close together; for soft material farther apart. The blades should be dipped into physiologic salt-solution to fill up the space between the blades. The instrument is then drawn through the tissue, and the section between the blades floated out in salt-solution. Some workers use this double-bladed knife to make a perpendicular cut into the tissue, then turning it to either side to cut the lower edge of the section and removing the blades with the section between them. Both the single- and the double-bladed razors require practice for successful section-cutting. Further, it must be borne in mind that sections of fresh tissue obtained by these methods are unsuited for complicated staining methods and can be used only for the simplest staining processes. Nevertheless, in a fairly large proportion of pathologic conditions it is possible to secure a diagnosis by these methods.

d. Freezing Microtome. Much more satisfactory sections can be obtained by freezing the fresh tissue and cutting it upon the freezing microtome. Various types of these instruments are in the market. The freezing is accomplished by the use of ether, ethyl-chloride or fluid carbonic acid gas. The ether and ethyl-chloride freezing-microtomes consist essentially of a metal plate or hollow box on which the tissue rests and against the under-surface of which a spray of ether or ethyl-chloride is forced by means of a rubber bulb connected with a supply of the freezing agent in a bottle. Such freezing attachments can be attached to any microtome, but special instruments as Jung’s “student’s freezing microtome,” Cathcart’s, Bausch and Lomb’s or Becker’s ether-freezing microtome can be recommended for this purpose. (See Fig. [52].) The tissue to be frozen must be of small size and not more than 3-4 mm. thick. It is placed upon the freezing plate in a drop of water, white of egg or thick gum-arabic, and pressed firmly against the plate. The spray must not be too constant or strong, but should be given with regular pauses of about a second to allow the ether to evaporate. The tissue must be firmly frozen, but not so hard that it crumbles. It is usually difficult to freeze the upper part of the tissue hard enough to give good sections, and this half-frozen tissue must be trimmed off with the knife until good sections are secured. As the sections are cut they are removed from the knife-blade with the finger and put into physiologic salt-solution that has been recently boiled to drive off the air, so that, in thawing, there may be no formation of air-bubbles in the tissue to cause artefacts. If put into strong alcohol diffusion-currents may damage the tissue; a succession of graded solutions should, therefore, be used if the sections are to be fixed and hardened. The further treatment of frozen sections is given below. The relative slowness of the method of freezing, its greater cost, the necessity of the frequent replacement of the rubber bulbs and tubing, and the greater amount of trouble required by the use of ether or ethyl-chloride are disadvantages that can be avoided by the use of the carbonic acid freezing microtome. Where much work is done by the freezing method the use of the latter is advised.

Fig. 52.—Type of freezing-microtome, for the use of ether or ethyl-chloride. Cathcart model.

Fig. 53.—Carbonic-acid freezing-microtome. Becker model.

Fig. 54.—Bardeen freezing-microtome attached to carbonic-acid gas-cylinder.

The carbonic acid outfit consists of a microtome arranged so that it can be connected with a cylinder of compressed carbonic acid gas, as shown in the Aschoff-Becker or Bardeen models. (Figs 53 and 54.) The cylinders or drums containing a charge of 15-30 lbs. of the liquid carbonic-acid are furnished by the trade at reasonable rates. It is not necessary to buy the drums, as they are replaced by full ones as needed. They are provided with valves and can be fastened in an upright position or laid flat upon a table, in any way convenient for the attachment to the microtome. Connection is made between the valve of the cylinder and the object-holder of the microtome by means of strong rubber tubing which should be securely wired at both ends, or by a flexible metallic tube. The latter is preferable, as the rubber-tubing often bursts, or is so stretched by the pressure of the gas that it must be frequently replaced. Object-holders with flexible metallic tube attachment for use in an ordinary microtome are supplied by Bausch and Lomb and other firms. My personal experience makes me prefer the Bardeen freezing microtome (see Fig. [54]) to all others. It is cheap and can be easily attached to the valve of the gas-cylinder, which is fastened horizontally to the table-top, as it has a screw-thread fitted to the uniform thread of the drum-valve. The object-holder can be raised or lowered between the glass-tracks on which the knife runs, so that sections of a definite thickness can be obtained. The most satisfactory knife is that of the type of a plane-bit set in a wooden handle. (See Fig. [55].) It must be well honed and stropped. The tissue, which should not be more than 5 mm. thick, is placed upon the object-holder in a drop of water, albumin fixative, or saturated solution of gum-arabic, and pressed firmly against the plate as the gas is turned on slowly and evenly. Freezing is usually accomplished in one-half to one minute. The best results are obtained by turning the gas on for about 15-20 seconds, then turning it off for several seconds; the tissue will continue to freeze; if not hard enough the gas is turned on again for 15 seconds and then turned off. If necessary this may be repeated until the tissue is properly frozen. The interruption of the gas-flow prevents over-freezing and thereby lessens the amount of change in the cells. When sufficiently frozen the gas is then turned off; the knife, with bevel-edge down, is held in the right hand with its handle between the palm and the ball of the thumb with the back of the hand uppermost; and the edge of the knife set at an angle of 45° to the tracks, along which it is rapidly pushed back and forth, shaving the sections from the frozen tissue as the latter is pushed up above the level of the tracks by means of the micrometer screw turned with the left hand. The screw should have been adjusted at the proper height before freezing, so that no time is lost in getting the object-holder up to the height for cutting. By holding the elbow and fore-arm closely against the body and pushing from the elbow with wrist-joint fixed the shaving of sections can be accomplished very quickly, so that with one freezing several hundred sections can be obtained. The microtome-screw should not be turned by the left hand until the knife on its return has cleared the tissue. A little practice in co-ordinating movements is necessary for expert work. The sections should be allowed to collect upon the knife-blade until a large number have been cut; they are then swept off the blade by the finger into cold freshly-boiled physiologic salt-solution. Sections of tissue that have been previously fixed can be put into 60 per cent alcohol, where the sections will unroll and straighten out perfectly flat. When the method of freezing by alternately turning on and off the gas, as given above, is followed, and the amount of freezing in the interval noted, there is no danger of over-freezing, and the breaking and crumbling of sections, with the production of marked artefacts, is avoided. When over-freezing has occurred the block may be partly thawed out by the finger. But over-freezing, as well as repeated freezing and thawing, may cause so much damage to fresh tissue that a diagnosis cannot be obtained from the frozen sections. After thawing out in the physiologic salt-solution the sections of fresh tissue obtained by freezing are treated according to the methods given in the next section of this chapter.

Fig. 55.—Knife for Bardeen freezing-microtome.

It must be emphasized here that the process of freezing is an active one, and alters the relation of cell-structures. With many fresh tissues the changes resulting from the freezing are so great that no diagnosis can be made. It seems necessary here to warn against the routine employment of the rapid method of freezing and staining fresh tissues in the diagnosis of material obtained by surgical operation. It has become a fad with some surgeons to make a pathologic diagnosis by the freezing method while the patient is on the table. Consequently, as the result of diagnoses made by the rapid freezing and staining method, many mistakes are made, even by supposed experts in this line. Particularly in the diagnosis of sarcoma is it easy to make mistakes because of the altered aspect of the cells caused by freezing. Normal lymphnodes, tonsils and inflammatory infiltrations may look like spindle-cell sarcoma in the sections prepared by the rapid freezing and staining method; and the exact nature of many other pathologic conditions cannot be accurately determined from such sections. On the other hand, a certain number of pathologic conditions can always be recognized in sections obtained in this way, and this fact justifies the employment of the method when properly controlled. In all cases in which the pathologic condition is not clearly evident in sections obtained by the rapid freezing and staining method no diagnosis should be given. In such cases the tissue should be fixed and then cut upon the freezing-microtome, or imbedded in celloidin or paraffin and then cut. Even when the patient is upon the table the tissue removed can be put into a 10 per cent formol solution for a few minutes and then frozen directly in gum without washing out the formalin. The longer the time that can be used for this short preliminary fixation the better the sections will be and the less the production of artefacts by the freezing. The process of fixation can be hastened by warming the fixing-fluid. I advise this short fixation before freezing in all cases of operative diagnostic work when the diagnosis is wanted as soon as possible. For all other work with the freezing microtome, when the question of time is not so important, fixation with formol for 12-24 hours should be carried out. This combination of formol-fixation and the freezing method permits the early diagnosis of autopsy and operation material, makes possible the demonstration of fat and other substances altered or dissolved out by the imbedding methods, and is a convenient way of selecting tissues, requiring more complicated staining processes. (See also Page [239].) The further treatment of sections of fresh tissue obtained by freezing will be found in the second section of this chapter.

8. Penciling or Shaking. For the demonstration of the stroma or reticulum either fresh or fixed sections may be placed upon a slide in an abundance of fluid and gently penciled with a fine, blunt camel’s hair brush until the fluid becomes cloudy. The cloudy fluid is washed away and replaced by fresh as long as cells are given off. The same results may be obtained by shaking the sections in a test-tube until the cells are shaken out of the stroma. The removal of the cells from the section is shown by its greater transparency. One of the practical applications of these methods in diagnostic work is the differentiation between alveolar round-cell sarcomata and carcinomata.

9. Digestion. For the demonstration of stroma, parasites, etc., the tissues may be digested with gastric or pancreatic ferments until the required elements are freed. A freshly prepared pepsin in 0.2 per cent HCl in the incubator for 3-5 hours will digest fibrin in fresh clots. Sections of fixed tissues may be imbedded in paraffin, cut, and digested on the slide with Grübler’s pancreatin according to the method of Flint.

10. Intravital and Supravital Staining. Various methods have been advised for the intravital staining of cell-granules. Intravenous or intraperitoneal injections of methylene blue, alum carmine, neutral red and other stains will produce intracellular granule-staining in various organs of experimental animals. In the study of low forms of animal life staining solutions may be injected, or the animal or its parts may be examined in staining fluids. Human material can be examined by this method immediately after being removed from the body by operation or within 1-2 hours after death. (For details of these methods see article on “Färbungen, intravitale,” Encyklopädie der mikroskopischen Technik.)

11. Injection. Injections for the demonstration of blood-vessels, lymphatics, ducts of glands, etc., are rarely used in pathologic work. The organs to be injected must be fresh, warm from the body, if possible. The vessels should be washed out by a freshly filtered 8 per cent sodium nitrate or sodium sulphate solution, followed by physiologic salt-solution. A cannula is introduced into a main vessel, tightly secured, and then connected with a syringe or a gravity injection-apparatus giving a constant pressure. The injection-mass is then injected under a low pressure. In the case of injections into lymphatic vessels the cannula should be introduced into the largest lymphatics at the periphery of the blood-vessels where larger lymph-vessels are more easily found. Injections are made with either cold or warm solutions; the latter are preferable but require that the organ to be injected be warmed by immersing it in water at a temperature of 40°C. The injection fluid must be of the same temperature. After the warm injection is given the organ is put into ice-cold 10 per cent formol solution until fixed and then after-hardened, imbedded and stained as desired. After the use of cold injections the tissues are fixed in 10 per cent formol, alcohol, or any other desired solution, and treated according to the end sought. Nuclear and diffuse stains contrasting with the color of the injection mass should be used. The process of injection requires great care; the pressure must be carefully regulated to prevent extravasations, and the injection-fluid must be free from air-bubbles. The injection is continued until the organ appears diffusely stained. Blood-vessels are fixed in their natural blood-injection by such agents as formol and chromic acid, so that stains acting upon the red blood cells cause the veins and capillaries to appear as if they had been injected.

The following injection masses are advised:—

1. Cold Injection Mass (Beale’s Glycerin Carmine):—

Dissolve 0.3 grm. of carmine in a small quantity of water containing 5 drops of ammonia; add 15 cc. of glycerin and shake; add drop by drop 15 cc. of glycerin containing 8-10 drops of glacial acetic acid. Then add further glycerin 15 cc., alcohol 8 cc., and water 24 cc.

2. Warm Injection Mass (Thiersch’s Berlin-blue Gelatin):—

(a) Dissolve 1 part of gelatin in 2 parts of water by allowing it to soak 24 hours, and then warming. Filter through flannel.

(b) Saturated water solution of ferrous sulphate.

(c) Saturated water solution of red ferricyanide of potassium.

(d) Saturated water solution of oxalic acid.

Make a solution (1) by adding 30 cc. of a to 12 cc. of b, and a solution (2) by adding 24 cc. of c to 60 cc. of a; both mixtures to be made at a temperature of 30°C. At same temperature add 24 cc. of d to solution 2, and then solution 1, stirring constantly so that Berlin blue is precipitated. Heat on a water-bath to 90°C.; filter through flannel.

3. Fischer’s Milk-Method. The vessels are flushed with 8 per cent sodium nitrate or sulphate solution, and then injected with milk. When sufficiently injected the tissue is hardened for 24 hours in a solution of water, 1,000 cc., formalin (40 per cent formaldehyde) 75 cc., and glacial acetic acid 15 cc. Freeze, cut and stain with Sudan III or Scharlach R.; the course of the vessels is outlined by the fat-globules.

4. Silbermann’s method of injecting indigo-carmine, eosin or phlosin-red into the circulating blood has been used for the demonstration of capillary thrombi, the latter remaining free from the pigment.

12. Warm Stage. For the study of vital phenomena in the living cell Ross’s electrical warmer is recommended. It can be slipped on and off the slide without changing the focus, and is managed without any difficulty. It keeps the centre of the slide at a temperature of 37°C. Reagents can be applied as desired. Deetjen’s agar may be used as a medium for the preservation of living cells. (See Methods of Blood-Examination.) Various forms of warm and moist chambers used in experimental embryological work can also be utilized in experimental pathology.

13. Tissue-cultivation. The embryologic methods of growing tissues in lymph and blood-plasma as developed by Harrison, Burrows and Carrel have been applied in pathologic work to the experimental study of repair and regeneration, grafting, transplantation and tumor-transplantation. (For methods see Harrison, Journal of Exper. Zoology, 1910; Burrows, Jour. of Amer. Med. Assoc., 1910; Carrel, Jour. of Amer. Med. Assoc., 1910.)

II. REAGENTS USED IN THE EXAMINATION OF FRESH TISSUES.

In the examination of fresh tissue it is often desirable to use certain reagents for the purpose of making chemical tests or to bring out some structures more prominently than others. To introduce these reagents beneath the cover-glass in such a way as to get the desired effect without disturbing given fields requires some practice with a very simple technical method. The preparation is first examined in salt-solution, and the cover-glass adjusted so that it has a slight rim of fluid about its edge, but not enough to make it float. The reagent to be applied is dropped with a glass-dropper at one side of the cover-glass, while at the other the salt-solution is removed slowly by a piece of absorbent paper. The changes produced in the tissue-elements during the progress of the reagent can be observed under low or high powers. Care must be taken to change the fluids so slowly that isolated cells will not be washed away.

1. Physiologic or Indifferent Fluids. Serous exudates, blood-serum, hydrocele fluid, etc.; artificial serum made by a mixture of 9 parts physiologic salt-solution with 1 part white of egg; or physiologic salt-solution (0.9 per cent for warm-blooded animals, 0.6 per cent for cold-blooded).

2. Maceration Fluids. 33 per cent alcohol (24 hours); chromic acid 1:5000 (24 hours); potassium bichromate 0.1-0.2 per cent solution (2-4 days for nervous tissue); 0.1 per cent osmic acid (12-24 hours); 33 per cent potassium hydroxide (¼-1 hour, for muscle, tissue must be examined in the solution, as the cells dissolve when water is added); Arnold’s iodine solution (10 parts of a 10 per cent potassium iodide solution to which are added 5-10 drops of a solution containing 5 grms. of iodine and 10 grms. of potassium iodide in 100 cc. of water. Macerate one or more days. If solution becomes discolored add more of the second solution); very dilute formol solutions (1 cc. to 500 cc. physiologic salt-solution); Müller’s fluid (2-3 days, good for nervous tissue).

3. Glycerin. Used without diluting as a clearing agent, particularly when pigment is present; and as a mounting medium for stained preparations that cannot be put into alcohol.

4. Potassium Acetate. Saturated water solution for clearing and mounting fresh preparations. Does not clear as strongly as glycerin, hence is better adapted for the examination of fresh tissues.

5. Acetic Acid. 1-2-5 per cent solutions are usually employed. Clears the protoplasm and causes the nucleus to shrink slightly and to stand out more distinctly. It differentiates fatty and albuminous granules, dissolving the latter; and is useful in the demonstration of elastic tissue fibres, sharply outlining these against the connective-tissue which swells and becomes clear.

6. Acetic Acid Fuchsin. A few drops of fuchsin are added to a 2 per cent solution of acetic acid. With this solution the nuclei are not only brought out more sharply but they are stained red.

7. Lugol’s Solution. Dilute Lugol’s solution to a pale yellow color. It brings out the contours of cell and nucleus, and has a specific reaction with glycogen and amyloid, giving both a brown color. Since glycogen is dissolved out in water-solutions tests for glycogen in fresh tissues should be made with iodine-glycerin or iodine-gum (Lugol’s one part, gum arabic 100 parts). Smears or cover-glass preparations may be placed in covered dishes containing a few crystals of iodine.

8. Potassium and Sodium Hydroxides. In solutions of 1-3 per cent all tissue-structures swell and dissolve or become unrecognizable except elastic fibres, fat, pigment, amyloid, bacteria, yeasts and moulds. Used especially for examination of skin-scrapings or pus for presence of blastomyces and various forms of parasitic moulds (barber’s itch, ringworm, tinea, etc.). Solutions of 33 per cent clear the tissues but do not destroy the cells. Useful for maceration. When diluted the cells are destroyed.

9. Mineral Acids. HCl or H2SO4 (3-5 per cent). Used to dissolve areas of calcification. Calcium dissolves with liberation of CO2; the phosphates dissolve without gas-formation, but with H2SO4 form crystals of calcium sulphate. Sulphuric acid is also used as a test for cholesterin (red or violet coloration), and with iodine as a test for amyloid.

10. Osmic Acid. In a 1 per cent solution this is used to test for the presence of fat (oleates), the fat-droplets become black or brown.

11. Sudan III or Scharlach R. Alcoholic solutions of these dyes are used for the demonstration of fat in fresh tissues. They stain fat orange to scarlet. (For method see Staining of Fat.)

12. Alcohol and Ether. Used to dissolve fat-granules and to differentiate between these and albuminous granules.

13. Stains. Fuchsin, methylene blue, methylene green in 1 per cent solutions in physiologic salt-solution and acetic-acid-fuchsin are the best stains used for the examination of fresh-tissues beneath the cover-glass. They are drawn under the cover-glass according to the method given above.

For the rapid staining of sections of fresh tissue cut by freezing a section is floated from the salt-solution onto a slide, which is then carefully lifted from the salt-solution and the excess of the latter removed. Several drops of methylene blue, carbol-thionin or carbol-kresyl-echt-violett are run upon the section with the glass-dropper, and allowed to remain for 15-30 seconds. The stain is then washed off with salt-solution, a cover-slip put on and the section examined in the salt-solution. Thionin has been especially recommended (Wood, Strouse and others) for the rapid staining of frozen sections; but I prefer to use carbol-kresyl-echt-violett (kresyl-echt-violett 1 grm., 5 per cent aqueous solution of phenol 80 cc., 95 per cent alcohol 20 cc.). This gives a very good differentiation in the section examined in water, and the picture is clearer than with thionin; the specific staining-reactions with mucin, amyloid, mast-cells, etc., are also more marked than with the latter stain. Such sections are not as clear as dehydrated and cleared sections and their possibilities of diagnosis are correspondingly limited, even in the hands of an expert with sections of this kind.

To make permanent balsam-mounts of the frozen unfixed sections the latter must be fixed in formol or alcohol, or by heat (hot water). The sections may be placed in 4 per cent formol for several minutes, then into 80 per cent alcohol, then stained, washed, dehydrated in absolute alcohol, cleared in carbol-xylol and finally mounted in balsam. Hæmatoxylin and eosin may be used for the staining. Fresh sections may also be fixed in hot water, stained in hæmatoxylin, dehydrated in alcohol and cleared in xylol. To save time the sections may be fixed on the slide for a few minutes in alcohol, care being taken to prevent the sections from rolling up by dropping the alcohol onto the middle of the section after it has been carefully flattened out on the slide. The section may then be stained with hæmatoxylin, borax-carmine or other stains, dehydrated, cleared and mounted in balsam. After fixing with alcohol on the slide the section may be attached to the slide by blotting it with absorbent paper, then covering section with absolute alcohol, draining this off after a few seconds and then running over the section a thin solution of celloidin, which is allowed to drain off, leaving a very thin film over the section and slide. The latter is then immersed in water for a few seconds, and the celloidin-film on setting holds the section to the slide, provided the celloidin has been of proper consistence. The section can now be stained, washed, dehydrated, cleared in origanum and mounted in balsam. When mounted the thin film of celloidin is invisible (Wright’s method).

I have originated a much better method which is in use in my laboratory, and can be applied to the staining of frozen sections of fresh tissues in large numbers for class use. The sections are floated from the salt-solution on to a warm solution of New Orleans baking molasses diluted ten times, or a dilute sugar-dextrin solution, and thence are floated on to a clean glass plate and arranged in rows. The plate is drained, and then without drying is immersed in absolute alcohol for 15-30 seconds; it is then flooded with a thin celloidin, drained, the celloidin film allowed to set, and the plate then put into warm water, where the celloidin sheet floats off, carrying the sections, which can now be cut out and treated as single celloidin sections, or the whole sheet can be carried through the staining, dehydrating and clearing solutions to be cut up into single sections before mounting.

CHAPTER XIX.
THE PRESERVATION OF MACROSCOPIC PREPARATIONS.

For preserving gross objects for museum specimens alcohol or formol may be employed. The former bleaches the tissues so that ultimately they are almost destitute of color. Formol in a 5 per cent solution gives better color-effects than alcohol, as the blood-containing parts remain darker. The fluid also remains clear and the tissues are firm. When alcohol is used the fluid must be frequently changed, as it becomes turbid and yellowish, and the tissues finally become soft and lose their form. The best methods of preserving the natural color are found in the various modifications of the Kaiserling method. The organs or tissues are placed first in a formol solution until they are just hardened, the formol changing the oxyhæmoglobin into acid hæmatin. They are then transferred to alcohol to bring back the natural color, which is accomplished by the change of the acid hæmatin to an alkali hæmatin, which has a color very closely resembling that of oxyhæmoglobin, so that the natural color is approximately reproduced. The method is carried out as follows:—

Sol. I.—Formalin200cc.
Water1,000cc.
Potassium nitrate15grms.
Potassium acetate30grms.

The tissues are left in this solution, in the dark, for one to several days, being watched carefully to see that they are not over-hardened.

Sol. II.—80 per cent alcohol for 1-6 hours and then 95 per cent until the color is fully restored (2-24 hours). Watch carefully and remove as soon as best color effect is reached, and preserve in—

Sol. III.—Glycerin400cc.
Water2,000cc.
Potassium acetate200grms.

The specimens must be kept in air-tight jars, and crystals of thymol added to prevent growth of moulds. This is sometimes very difficult, and it becomes necessary to change the discolored fluid for clear. I have found the rectangular museum jars best adapted for the preservation of Kaiserling specimens. I use a wooden top which fits over a thick piece of felt cut just the size of the jar, which in turn fits over a piece of dental rubber cut to fit the jar. The jar is placed upon a wooden bottom which has upright steel rods at the corners, that pass through holes in the wooden top, and have a screw-thread so that they can be fitted with screws, which when screwed down hold the wooden top, felt and rubber sheeting tightly in place, making the jar air-tight, but giving a top easily removable. Very beautiful specimens can be secured by the Kaiserling method, and they can be kept for several years, but sooner or later the color-effect is lost. Light, heat and exposure to the air cause a loss of color.

Some workers prefer the following in place of Sol. I:—

Hot water2,000cc.
Sodium sulphate40grms.
Magnesium sulphate40grms.
Sodium chloride20grms.

When salts are dissolved and solution cool add 200 cc. of formalin.

Melnikow-Raswedenkow Method:—

Sol. I.—Water100parts
Formol10parts
Sodium acetate3parts
Potassium chlorate0.5part

Leave in this 1-2-3-4-5 days, according to size of specimen. Large organs must have solution injected into vessels.

Sol. II.— 95 per cent alcohol, until color is restored.

Sol. III.—Preserve in: Water100parts
Glycerin60parts
Potassium acetate30parts

Pick’s Method:—

Sol. I.—Water1,000cc.
Formol50cc.
Carlsbad salts50grms.

Then transfer to 80 and 95 per cent alcohols, as for Kaiserling, and preserve in water 9,000 cc., glycerin 5,400 cc., sodium acetate 2,700 grms.

Westenhoeffer’s method of preserving uric-acid: formol vapor 4-24 hours, then 80-90 per cent alcohol containing mercuric oxide, and preserve in glycerin to which some mercuric oxide covered by cotton or absorbent paper has been added.

Claudius’s Method. The specimen is placed on a grating in a closed vessel containing a concentrated solution of ammonium sulphate, an abundance of the crystals being left on the bottom of the vessel. Carbonic acid or illuminating gas is passed through the ammonium sulphate solution for 48-72 hours, and the specimens are then preserved in the same solution. My experience with this method has not been satisfactory.

Gelatin method of mounting Kaiserling preparations: Soak washed Gold Label gelatin in distilled water for 12-24 hours. Take equal parts of water-logged gelatin and glycerin and dissolve by heating in a double boiler, stirring, for 15-20 minutes. Cool to 40°C. Then clarify with white of egg (well-whipped whites of three eggs to half a gallon of jelly: stir well; steam for ½ hour) and filter through cotton-wool. Add to jelly a few drops of a weak aqueous solution of crystal violet to remove yellow color (Bruère and Kaufmann). To prevent growth of bacteria a small percentage of formol or crystal of phenol may be added. Kaiserling specimens are placed in glass dishes in the melted jelly, and covered with it. When set the dishes containing the mounted specimens may be covered with glass-plates and fastened to these by balsam or cement. I use a deep Petri dish, filling it about two-thirds full with the jelly; over this I pour melted paraffin of a very low melting point so as not to melt the jelly. When the layer of soft paraffin is hard, a thick layer of paraffin of a 52° melting-point is poured over it, and the dish filled even. When the hard paraffin is set, it is varnished with shellac. Liquefaction of the gelatin by bacteria or enzymes constitutes the great drawback to this method.

CHAPTER XX.
THE FIXATION AND HARDENING OF TISSUES.

GENERAL CONSIDERATIONS. For the examination of material that may be injured by freezing, or when very thin sections are required for complicated staining procedures, it becomes necessary to prepare the tissue by fixation and hardening, so that it can be imbedded in some medium permitting the cutting on the microtome of as thin sections as may be desired. Fixation is that process by which the appearances of the tissue are preserved as they were when it was taken for examination; hence in order to obtain pictures resembling as closely as possible those of the living tissue the material should be fixed immediately upon its removal from the living body by operation, or as soon as possible after death when obtained by autopsy. Fixing agents act by coagulating the cell albumins, in this way “setting” or “fixing” the constituents of the cell so that further change is stopped. Fixation, therefore, hardens the cell, and all fixatives are also hardening agents. A practical distinction between fixing and hardening is made, however, resting upon the fact that not all fixatives harden the tissue so completely that the proper consistence for the cutting of thin sections is attained. To achieve this the tissue must be dehydrated. Alcohol and acetone are the only reagents fixing and hardening perfectly at the same time, as they remove the water from the tissue; for all other fixing agents an after-hardening in alcohol is necessary. In the case of such reagents the division of the process into a primary fixation and a second hardening stage has been the cause of the divergence in meaning of the two terms.

The best fixing agents are those that kill the cells at once, but cause a slow coagulation with little or no shrinking. They must penetrate and diffuse through the tissues rapidly so that the deepest cells are quickly reached. Acid media, especially those containing small percentages of acetic acid, are therefore better than alkaline solutions. The tissue-elements, particularly the nuclei, must be preserved as perfectly as possible so that they will not be affected by further procedures of microscopic technique. The chemic properties of physiologic and pathologic substances must likewise be preserved. The preservation of karyokinetic figures is a criterion of good fixation. In pathologic work it is also desirable that the fixing agent should preserve the red blood cells, and permit of the staining of bacteria in sections. Since fixing media are more or less selective in their action, it follows that there is no one fixative that gives equally good results in all cases. Especial fixing reagents must be used for the demonstration of certain substances (fat, etc.), or for the use of certain staining methods. Some stains cannot be used at all after certain fixing agents have been employed. For general use that fixing agent having the widest range of usefulness should be employed; and for this reason fixing media composed of several fixing agents are often employed in preference to the use of a single one.

GENERAL RULES FOR FIXING AND HARDENING.

The tissue should be put into the fixing fluid as soon as possible after its removal from the body. It must not be allowed to dry. There should be an abundance of the fixing solution, 25-50 times the volume of the object to be fixed. The tissue should never be put into a dry vessel and the fixing fluid poured upon it; the vessel should first be filled with the fixing solution and the tissue then dropped into the latter. A slight agitation of the fluid will prevent the sticking of the tissue to the bottom or sides of the vessel. The size of the pieces must be adapted to the penetrating power of the fixing reagent, but as a rule the pieces should not be more than 2-3 cms. in thickness, and for some reagents 0.5 cm. is as thick as they can safely be. The reagents used should be changed when they become cloudy or discolored. The used solution may be filtered and used again, but some reagents can be used but once. Alcohol may be saved for redistillation. In the case of some reagents (mercuric chloride, chromic acid, osmic acid, etc.), the time limits of the fixation should not be exceeded, as over-fixation will ruin the staining-power of the tissues. Alcohol is practically the only solution in which tissues may be left indefinitely, but even with it there are certain limitations. As a general rule the time required for fixation may be shortened by keeping the reagents at incubator-temperature. As the different fixing reagents vary so greatly as to their especial advantages and disadvantages these will be considered separately. Only the best and most commonly used methods are here included.

1. ACETONE. A water-free acetone is employed by placing pure dried white copper sulphate in the bottom of the bottle or vessel in which the fixation-process is carried on. Several layers of filter-paper are put over the copper sulphate to keep the tissues from touching it, and the acetone is then poured into the vessel. As soon as the copper-sulphate becomes blue it must be again fused. It is only by this method of constant dehydration that acetone can be employed to any advantage as a fixing agent; if fused copper sulphate is not employed the amount of acetone necessary to fix well is so great that the method becomes too troublesome and expensive to be recommended. But with the simple copper-sulphate method of constant dehydration acetone becomes the cheapest, most rapid and one of the best fixing reagents. The use of alcohol is avoided, and the period of infiltration in xylol and in paraffin shortened. For very quick work the entire process of fixation, hardening and dehydration may be achieved by the use of acetone alone, small pieces of tissue being fixed ½-2 hours in acetone and then transferred directly to soft paraffin. For ordinary work pieces of tissue 0.5 cm. thick are put into acetone over fused copper sulphate for 20-60 minutes. A judgment of the degree of fixation can be obtained by pressing the tissue lightly between the fingers; if it is of uniform consistence, and does not give as if the inner portions were softer than the surface, the fixation is complete. From the acetone the tissues may be brought directly into xylol for 5-10 minutes, until they obtain a cloudy transparency, thence into paraffin for 15-30 minutes and then blocked. The whole process of fixation, imbedding, cutting and staining can be carried out in 30 minutes. Acetone may be combined with formol, alcohol or any of the other fixing agents, but when it is desired to use any one of these for some especial purpose it is better to fix first with the desired reagent and then to use acetone for the dehydration-process alone, instead of alcohol. We have found that formol-fixation followed by acetone dehydration gives excellent results for general pathologic work. Formol-acetone (acetone 100 cc., formol 10 cc.) may be found to have advantages. The quick fixation in water-free acetone causes less contraction than fixation with absolute alcohol; fixation with graded acetone-solutions causes practically none.

Advantages. It is the cheapest and quickest method. It penetrates well and causes little contraction of the tissues, shortens the time in xylol and paraffin and makes more easy the cutting of dense fibrous structures. Cell-division figures are as well preserved as by alcohol fixation, and the staining of bacteria in the tissues can be carried out as well after acetone-fixation as after alcohol. It preserves lecithin, hence can be used for the fixation of nerve-tissues. When combined with formol (formol-acetone) the red blood-cells are well-preserved and take a brilliant eosin stain.

Disadvantages. The disadvantages are practically the same as with alcohol fixation but not so marked. Fat is dissolved, cell-division figures are not so well-preserved as with mercuric chloride and Flemming’s solution, and the blood-cells not so well preserved as with formol and mercuric chloride.

2. ALCOHOL. Absolute, 95-96 per cent alcohol, or graded alcohols (70, 95 per cent. and absolute) may be used for fixation. For this purpose the stronger alcohols are preferable, as the weaker solutions do not fix quickly enough. On the whole the use of 95-96 per cent is to be advised. The pieces of tissue must not be thick. Plenty of alcohol should be used and it should be changed several times during the process of fixation, which for larger pieces requires several days. For the last change absolute should be used. Very small bits such as uterine curettings can be fixed in one hour, by using three changes of absolute alcohol. Since alcohol both fixes and hardens it has been generally used, but it is a relatively poor fixative. For after-hardening it is indispensable. Absolute alcohol may be made from 96 per cent by the use of fused copper sulphate. To test the strength of alcohol mix a few drops with pure water-free xylol; if no sediment appears when viewed against a dark background the alcohol is absolute or practically so. Many of the disadvantages of alcohol fixation can be obviated by the use of formol-alcohol (95 per cent alcohol 100 cc., formalin 10 cc.).

Advantages. Cheapness, quickness, and ease of method. Can be used for quick diagnostic work. Penetrates well, and can be used for large pieces of tissue. Preserves glycogen, is especially good for the staining of bacteria in sections, and the majority of stains work well with alcohol-fixation.

Disadvantages. Causes much shrinking and loss of finer details; preserves cell-division figures not at all or poorly; destroys the red blood cells; dissolves fat and other chemic products: does not permit of the use of certain specific staining methods (nerve-tissues); causes excessive hardness of fibrous and elastic tissues and makes cutting difficult.

3. CHROMIC ACID AND SALTS. Chromic acid is rarely used alone in pathologic work, but is a constituent of Flemming’s solution (see below). Its salts are employed in the form of:—

A. Müller’s Fluid (Potassium bichromate 25.0 grms., sodium sulphate 10.0 grms., water 1,000.0 cc.) This was formerly the favorite fixing solution, but is now used chiefly for the eye and nervous tissues, either alone, or after formol fixation, or mixed with formol. Large pieces may be used, even an entire brain, but the process requires months or even a year for the best results. Even small pieces take several weeks. The process may be hastened in the incubator. The solution should be changed whenever it becomes cloudy. When fixation is complete the fixed and hardened tissue may be cut directly on the freezing-microtome or after-hardened and dehydrated in alcohol or acetone when it is to be imbedded. The dehydration in alcohol should be carried out in the dark and without previous washing of the tissue in the case of nervous tissue. Greenish or brownish chrome precipitates appear if the dehydration takes place in the light; but for ordinary material this precipitate can be avoided by washing the fixed material in running water for 24 hours before transferring to alcohol. Moulds grow luxuriantly in Müller’s fluid, but may be inhibited by the use of pieces of camphor, thymol, etc.

Advantages. Cheap, penetrates well, causes little shrinking, permits special nerve-stains, preserves red blood-cells, gives beautiful results with ordinary stains, preserves fat.

Disadvantages. Slowness; does not preserve division-figures; does not permit of staining for bacteria; does not give good results with many special staining methods (fibrin, elastic tissue, reticulum, etc.).

B. Erlitzky’s Fluid (Potassium bichromate 25.0 grms., copper sulphate 5.0 grms., water 1,000.0 cc.). Used for fixation of nervous tissue. The formation of pigment precipitates may lead to misinterpretation: the artefacts may be removed by hot water or dilute acetic acid.

C. Orth’s Fluid (Müller’s fluid 100 cc., formol 10 cc. Make fresh before using, as the mixture precipitates on standing.). Fix for 3-12 hours in the incubator, or for 24-48 hours at room temperature. Wash in running water for 12-24 hours; cut on the freezing microtome, or after-harden in acetone or alcohol and imbed.

Advantages. Combines the good features of Müller’s and formol fixations, and obviates some of the disadvantages. It is a good general fixing solution.

3. FORMOL OR FORMALIN (40 per cent solution of formaldehyde gas). Used in a ten per cent solution (water or physiologic salt solution nine parts, formalin one part), often incorrectly called 4 per cent formol or formalin, the mistake arising from the confusion with 4 per cent formaldehyde gas. For nerve-tissues it is better to dilute the formol with physiologic salt-solution than with water. Fix for 3-4-12 hours according to size of tissue. As it penetrates well large pieces can be used. Wash in water before after-hardening in alcohol, if tissue is to be employed for general work, otherwise it can be transferred directly to the alcohol without washing. Tissues can be kept in formol for some weeks, but after that time the staining-power is slowly affected, and the finer structures suffer.

Advantages. Probably the best fixing reagent for general pathologic work. It is cheap; easily made and kept in solutions of proper strength for fixing; hardens while it fixes; does not require after-washing; penetrates well; causes little shrinking; permits freezing directly from the fixing solution; preserves fat; permits the use of after-hardening with bichromate solutions for especial nerve-stains: preserves the red blood-cells: gives good results with nearly all stains, and differentiates bile-pigment from hæmatoidin. It is the best fixing reagent when tissues are to be sent some distance, as over-fixation occurs only after several weeks or even months.

Disadvantages. Affects many people unpleasantly, causing coryza, eczema of the hands and arms, and affections of the finger-nails, so that workers having acquired this idiosyncrasy cannot expose themselves to formol vapor; it dissolves glycogen and uric acid; does not fix cell-division figures as well as mercuric chloride or Flemming’s solution; causes the precipitation of diffuse hæmoglobin in the form of brown or black pigment-granules that may be taken for melanin, malaria pigment or hæmosiderin; causes a pseudo-ochronosis of cartilages; and, unless thoroughly washed from the tissues before after-hardening in alcohol, it makes carmine-staining difficult or unsatisfactory and affects also the specific staining-reactions for amyloid and mucin; it is not as good as alcohol or mercuric-chloride when the sections are to be stained for bacteria. In spite of these disadvantages it can be recommended as the best general fixing reagent.

For Orth’s fluid, formol-acetone and formol-alcohol see above.

4. FREEZING AND DRYING. The fresh tissue is frozen and dehydrated in a vacuum over sulphuric acid, at a temperature of 20-30°C.; when completely dried it is imbedded directly in paraffin. This method has been especially recommended by Altmann on the ground that the tissues are simply deprived of water without any change in volume.

5. HEAT. Physiologic salt-solution is heated to 80°C. Thin pieces of tissue are placed in the hot water for two minutes, and then after-hardened in alcohol. For larger pieces of œdematous tissues, cysts, etc., that cannot be cut into thin pieces, the salt-solution should be brought to 100°C and the tissues boiled for several minutes. This method is advised particularly for the coagulation of albumin in cysts, œdematous tissues, for the study of renal casts, etc.

6. MERCURIC CHLORIDE. This is used most commonly in the form of a concentrated water solution (mercuric chloride 7.5 grms., sodium chloride 0.5 grm., glacial acetic acid 5 cc., water 100.0 cc.), or as Zenker’s solution (mercuric chloride 5.0 grms., sodium sulphate 1.0 grm., potassium bichromate 2.5 grms., water 100 cc.; dissolve by heating, add 5 cc. glacial acetic acid just before using. The use of a 5 per cent formol solution instead of acetic acid is recommended). The pieces of tissue should not be thicker than 5 mm. Fix 6-24 hours, then wash 24 hours in running water, and after-harden in alcohol. Should the sections show mercuric precipitates they should be treated with Lugol’s solution for 30-60 minutes, then washed in a dilute solution of lithium carbonate and thoroughly washed out in water and alcohol. Much better stains can be obtained by this treatment of the sections with Lugol’s. The use of iodine in the alcohol during the process of after-hardening is not advisable because of the action of the iodine upon the albuminates of mercury. A 5 per cent solution of sublamine in distilled water has recently been recommended by Klingmüller and Veiel. Fix 1-3 hours, wash and after-harden in alcohol. Precipitates are not formed, and good staining-results are obtained.

Advantages. The mercuric chloride solutions preserve well the red blood-cells, mitotic figures and finer details of cell-structure, and permit the staining of bacteria and animal parasites in the sections. Certain especial staining methods (Mallory’s reticulum-stain, etc.) can be used only after mercuric chloride fixation, while many others (Heidenhain’s iron-hæmatoxylin, Biondi-Heidenhain triple stain, etc.) give best results after this fixation. For ordinary work the saturated mercuric chloride solution is preferable to Zenker’s, as the latter does not give good results with the commonly-used hæmatoxylin stains.

Disadvantages. More troublesome and expensive; require thorough washing and subsequent removal of precipitates, and affect (Zenker’s particularly) certain stains.

7. OSMIC ACID. Osmic acid is used alone in a 1 per cent solution, or in such combinations as Flemming’s solution (chromic acid 1 per cent. sol. 15 cc., 1 per cent osmic acid 4 cc., glacial acetic acid 1 cc.), Hermann’s solution (same as Flemming’s, with 15 cc. of a 1 per cent platinic chloride substituted for the chromic acid), Altmann’s solution (5 per cent potassium bichromate solution 50 cc., 2 per cent osmic acid solution 50 cc.), Marchi’s solution (Müller’s fluid 2 parts, 1 per cent osmic acid solution 1 part), and that of Pianese (1 per cent sodium-chloroplatinate 15 cc., 2 per cent osmic acid 5 cc., ¼ per cent chromic acid 5 cc., formic acid 1 drop). The pieces of tissue must be very thin, as osmic acid penetrates very slightly. Fix 6-24 hours in the dark, and wash thoroughly in running water, and after-harden in graded alcohols. These solutions have but limited use in pathology, and are used chiefly for the study of fat (oleates) and mitotic figures. Flemming’s and Hermann’s solutions are the best for the study of mitotic figures, the latter bringing out plasma details more clearly. Marchi’s fluid is used especially for the study of nerve-degeneration, and Altmann’s for the demonstration of Altmann’s granules. The method of Pianese is used for the demonstration of cell-inclusions. The osmic-acid mixtures are all expensive, penetrate poorly, cause precipitates, and affect greatly the staining-power of the tissues, so that it becomes necessary to use certain stains (safranin, carbol fuchsin, aniline gentian violet, etc.) as counterstains.

8. PICRIC ACID. A saturated water solution of picric acid is usually employed. Fix 12-24 hours, and wash in alcohol, not water, and after-harden in graded alcohols. Preserves mitotic figures, fine details of cell-structure, and is very good for bone and calcified tissues, as it decalcifies and fixes at the same time.

Numerous modifications and combinations of the above methods have been proposed such as Flemming’s chrom-acetic solution, Rawitz’s chromic-picric-nitric fluid, Rabl’s chrom-formic mixture, Burckhardt’s chrom-osmic-nitric solution, Merkel’s fluid (chromic acid-platinic chloride), Carnoy’s mixture (glacial acetic acid 1, absolute alcohol 6, chloroform 3), and many others. They have a limited use in pathologic work.

In the judgment of a section as to its fixation the following points may be of service: in alcohol fixation the red blood cells are haemolyzed, and there is much shrinking; with formol fixation the red cells stain copper-red with eosin; in mercuric chloride fixations the red cells stain rose-red with eosin, and pigment precipitates are present; in bichromate fixation the red cells preserve their natural color, and fat cells show a brownish color; osmic-acid fixation is shown by the black color of the oleates, and the failure of the tissue to stain by ordinary stains.

CHAPTER XXI.
DECALCIFICATION.

Bone and tissue containing deposits of lime must be decalcified before they can be sectioned on the microtome. The decalcification should be carried out after fixation and before the after-hardening in alcohol. Some reagents may combine decalcification with fixation, but this is satisfactory only when the amount of lime-salts is relatively small. Fresh tissues should not be put into any of the stronger acid decalcifying fluids, as they alter unfixed cells so that the staining-power is lost and the fine histologic details destroyed. The fixed tissue cut into small pieces is put into the decalcifying reagent, which is used in large amount and must be frequently changed. It is left in the decalcifying fluid until the calcium salts are removed, as shown by tests with needle or scalpel. The tissue must not be left in the fluid after decalcification is attained, as the staining-power is affected by all decalcifying reagents; it is therefore necessary to make frequent tests in order to judge of the progress of the decalcifying process. After decalcification the tissue should be washed in running water for 24 hours, and then after-hardened in alcohol. Alkaline solutions may be used to remove the acid before washing. Sections of decalcified tissue always stain slowly, and it is advisable to remove any acid remaining in the tissues by soaking the sections in a saturated water solution of lithium carbonate before staining. Numerous formulæ for decalcifying fluids have been recommended; a few of the best methods only are given here.

1. Combined Fixation and Decalcification. Picric acid or Müller’s fluid may be used for this purpose when the amount of lime-salts contained in the tissues is very small. The process is slow.

2. Trichloracetic Acid. Fix tissues in 10% formalin and decalcify in trichloracetic acid 90 cc., formol (40 per cent formaldehyde) 10 cc. Change frequently. Decalcification is rapid, the tissue is but little changed and the staining-power not affected.

3. Concentrated Sulphurous Acid. Fix in 10 per cent formol; decalcify in concentrated sulphurous acid for 24 hours or longer if necessary. Wash thoroughly in alkaline water. This is a very good and rapid method; the staining-power is but little affected.

4. Haug’s Solution. (Pure nitric acid 3-9 cc., absolute alcohol 70 cc., sodium chloride 0.25 grm., water 30 cc.). For tissues fixed in mercuric chloride.

5. Phloroglucin. (Phloroglucin 1 grm., pure nitric acid 10 cc., distilled water 50 cc.). The solution must be carefully dissolved over the flame in a hood. Decalcification is rapid, and the tissue is protected from the acid by the phloroglucin.

6. Ebner’s Fluid. (Hydrochloric acid 5 cc., sodium sulphate 5 grms., alcohol 500 cc., water 1,000 cc.).

7. Schaffer’s Method. Imbed the fixed and hardened tissue in celloidin, harden the celloidin preparation in 85 per cent alcohol, then place celloidin block in a 3-5 per cent water solution of pure nitric acid and agitate in Thoma’s water wheel, for 12 hours, or longer according to the size of the piece. Transfer block to a 5 per cent solution of lithium carbonate or sodium sulphate for 12-24 hours, changing solution several times, wash in running water for 48 hours, dehydrate in graded alcohols up to 85 per cent, and cut.

CHAPTER XXII.
IMBEDDING.

The most perfect methods of fixation and hardening do not permit the cutting of fine sections on a microtome without the freezing of the tissue, or its infiltration and imbedding in some substance which surrounds it with a protective coating, and preserves and holds together its structural elements in their relative positions. For the cutting of very thin sections, or for the preparation of serial sections, it is absolutely necessary to employ the process of imbedding. At the present time paraffin and celloidin are the two substances in general use for this purpose. While each one of these possesses certain advantages over the other, and we find consequently one laboratory worker preferring celloidin and another paraffin for general work, a long and varied experience makes me believe that for a teaching laboratory and for diagnostic work when much material is examined, the paraffin method answers all purposes much better than the celloidin, and that the latter need be employed only in very exceptional cases. Since paraffin sections can be transferred into celloidin by the molasses- or dextrin-fixative method, thus enabling the use of staining-methods that require celloidin sections, very few advantages are left in the favor of celloidin as an imbedding agent. The paraffin method requires a more expensive outfit to start with in the form of a paraffin oven and thermo-regulator, but otherwise the two methods cost about the same. The paraffin method requires more careful attention than the celloidin. As a rule thinner sections can be obtained in paraffin than in celloidin, and for the preparation of serial sections the paraffin method is the only method. Paraffin blocks can be labeled and filed away, and kept indefinitely without any loss of staining-power. With careful attention paid to the different steps of the imbedding process practically everything that can be cut in celloidin can be cut in paraffin. For very large pieces a slow imbedding in celloidin is, however, preferred by most workers. Hard and brittle tissues are as a rule more easily cut in celloidin. For the staining of bacteria in sections paraffin imbedding is necessary. Both methods should be learned and practiced with equal facility; a working knowledge of both is essential in pathologic investigation and diagnosis.

1. CELLOIDIN IMBEDDING. The granular form of Schering’s celloidin is the best preparation to use, although good results can be obtained by using a cheaper well-washed gun-cotton. In purchasing the latter care should be taken to secure a sample that dissolves easily in alcohol and ether, and does not give off yellow fumes when exposed to the light. Schering’s granular celloidin keeps well, and forms on solution a firm, tough, transparent imbedding mass, so that thin sections are obtainable without difficulty. When kept long in stock celloidin becomes hard and dissolves more slowly. For use three solutions are made, thick (10 per cent), thin (2 per cent), and medium (5 per cent). The celloidin granules or shavings are put into a wide-mouthed bottle having a tight stopper, and are covered with absolute alcohol and well shaken, and left for 24 hours. An equal quantity of pure ether is then added, the mixture is well stirred and allowed to stand for another 24 hours, when it is again stirred and evenly mixed, and is then ready for use. When gun-cotton is used it is torn into fine shreds and added to a mixture of equal parts of absolute alcohol and pure ether and shaken until sufficient has been added to give the solution the desired strength.

Slow Celloidin Method.

1. Absolute alcohol 24 hours.
2. Equal parts absolute alcohol and ether 24 hours.
3. Thin celloidin for 1-3 days.
4. Medium celloidin for 1-3 days.
5. Thick celloidin for 1-3 days.
6. Block.

The tissue is blocked by removing it from the thick celloidin on a section-lifter and placing it on a block of vulcanized fiber or wood with enough of the thick celloidin about it to form a good matrix. The preparation is then allowed to evaporate in the air until the surface of the celloidin becomes firm (does not stick to the finger). The block is then placed in 80 per cent alcohol or pure chloroform until hard enough for cutting (1-24 hours). Cork should not be used for blocking, nor should wood unless the tannin has been removed by long treatment with alcohol-ether. The celloidin will adhere more firmly to the fiber block if the latter is dry, and if there is a sufficient layer of celloidin between the tissue and the block. The imbedded tissues may be preserved in 80 per cent alcohol for a long time, but gradually lose their staining power. They may also be kept dry by coating them with melted paraffin. The blocks when preserved in alcohol may be marked with an indelible pencil.

For imbedding large pieces of tissue in celloidin a glass dish may be filled with thick celloidin and the infiltrated tissue placed in it with cutting surface down. The celloidin is then allowed to evaporate slowly under a glass cover, and fresh celloidin may be added as shrinkage occurs. When well-hardened the celloidin is cut out of the dish and the block trimmed to the proper proportions, and attached directly to the object-holder of the microtome or to a block of wood by a few drops of thick celloidin, allowing it to dry for a minute or so and then immersing in 80 per cent alcohol. The block may be cut on the freezing microtome by soaking the hardened celloidin in water to remove the alcohol (when block sinks), then coating it with saturated gum arabic solution, and freezing.

Rapid Celloidin Method.

I have used the following method in my laboratory for a number of years as a regular procedure in practical diagnostic work, and the results have been uniformly good.

1. Fresh tissue cut thin, or uterine curettings, in absolute alcohol 1½ hours, three changes of fresh absolute during this time.

2. Alcohol-ether 1 hour.

3. Thin celloidin at incubator temperature 12 hours (over night).

4. Medium celloidin 1 hour.

5. Block from medium celloidin, evaporating celloidin by blowing, and building up matrix by adding successive layers of celloidin.

6. 80 per cent alcohol 1-3 hours.

7. Cut.

The quick celloidin methods recommended in the literature (Kaufmann, Stepanow, Scholz and others) require more time, and do not give better results. Material received from operative clinics late in the afternoon can be sectioned and stained usually by ten o’clock the next morning.

2. PARAFFIN IMBEDDING. A paraffin of a melting-point sufficiently high enough to withstand summer heat is advisable; a 52°C. paraffin answers for this latitude. The use of softer paraffins is not necessary. The paraffin-oven should be regulated at a constant temperature of 54-55°C. Over-heating of the tissue while in the oven must be carefully safeguarded, so that in the management of a paraffin-oven the care of the thermo-regulator is the most important thing.

Slow Paraffin Method.

1. Thorough dehydration in absolute alcohol 12-24 hours.
2. Aniline oil to remove alcohol, until tissue becomes transparent or sinks.
3. 1st. Xylol, ½ hour, to remove aniline oil.
4. 2nd. Xylol, 1-2 hours, until translucent.
5. 1st. Paraffin, 52°C. in oven, 1 hour, to remove xylol.
6. 2nd. Paraffin, 52°C. in oven, 1-12 hours.
7. Block.

The use of xylol-paraffin is not necessary. For blocking staining dishes, watch-glasses, glass salt-cellars, paper-boxes, metal frames., etc., may be used. A thin smear of tincture of green soap or glycerin is rubbed over the inside of the imbedding box, and it is nearly filled with fresh melted paraffin. With warm forceps the tissue is taken out of the bottle of second paraffin in the oven, and arranged in the melted paraffin in the imbedding dish in the proper position for cutting. Care must be taken that the melted paraffin is not hot enough to “burn” the tissue, else its staining-power may be affected. The surface of the paraffin is then cooled by blowing upon it, and as soon as a film appears upon the surface, the dish is carefully immersed in cold water, so that the paraffin may set quickly. When cool the paraffin-block should slip out of the dish and float to the surface. It is then trimmed to the desired shape, leaving a good matrix of paraffin around the tissue. The paraffin-block is then fastened to a wooden block or to the object-holder of the microtome by means of melted paraffin (a hot knife is drawn along the under-surface of the block and the latter immediately pressed upon the wooden block or object-holder). Chloroform, cedar oil, benzene, carbon bisulphide, etc., may be used instead of xylol, and each one possesses certain advantages for certain purposes. Benzene is advisable for osmic-acid preparations.

Rapid Paraffin Method.

The above method can be greatly shortened for uterine curettings, thin bits of tissue, etc., if the various steps are closely watched, and if the entire process is carried on in the oven. The whole process may be carried out in 1-3 hours, a very great advantage over the quick celloidin method. A simpler and cheaper method is that recommended by Heller, Henke and Brunk, as follows:—

Acetone Method.

1. Small bits of fresh tissue, or tissue fixed in formol, in water-free acetone over copper sulphate for ½-1½ hours.

2. Transfer tissue directly to fluid paraffin in the oven for ½-1½ hours; the acetone evaporates, and the tissue is infiltrated with paraffin; or put into xylol 5-10 minutes, then in paraffin 15-20 minutes.

3. Block.

By this method the entire process of fixing, hardening, imbedding, cutting and staining can be carried out in half an hour, and by it the freezing-microtome can be dispensed with in a large part of quick diagnostic work.

Pyridin Method.

1. Fix in formol.

2. Dehydrate and clear in pyridin.

3. Paraffin.

This method requires a longer time than the acetone method, and is not so good.

Combinations of celloidin and paraffin may be employed by imbedding first in celloidin, transferring block to origanum oil, then xylol and finally paraffin. Formol-agar has been recommended by Bolton and Harris for simultaneous fixation and imbedding. It offers no especial advantages. Wright uses formol-gelatin for imbedding tissues for sectioning on the freezing microtome. The bits of tissue are placed in warm 20 per cent pure gelatin; this is allowed to set, and the block placed in 10 per cent formol for 24 hours; it is then frozen and cut. Soap-gelatin, glycerin-gelatin, gum-glycerin, etc., are now rarely used for imbedding.

Note:—When a number of pieces of tissue are blocked at the same time, either in celloidin or paraffin, they may be tagged with paper labels fastened to the tissue with a drop of gum tragacanth or gum arabic. As the gum is not soluble in any of the infiltrating and imbedding media the labels remain attached until the block is ready for cutting.

CHAPTER XXIII.
SECTION-CUTTING.

Fixed and hardened tissues may be sectioned on a microtome without freezing or imbedding, or they may be cut on a freezing-microtome; or, having been imbedded in either celloidin or paraffin, they may be sectioned on any form of microtome suited to the purpose desired. The choice of the microtome depends, therefore, upon the character of the work. For ordinary purposes the small sliding microtomes furnished with a crank are preferable, as they are more easily kept in order and can be used for rapid cutting. For serial paraffin sections the Minot automatic rotary microtome is advised. Especial types of microtomes can be obtained for the cutting of large sections, particularly for brain-sections. The Bardeen freezing-microtome I have already recommended as the most practical instrument for freezing work. In all cases it is absolutely necessary that the instruments be in good order and that they work with precision. The microtome knife must be carefully honed and stropped. A heavy, rigid, biconcave knife-blade should be used, and a good hone and strop are absolute necessities. In honing, the knife is drawn from heel to toe with cutting edge forward; while, in stropping, the motion is reversed, the back of the knife being forward, and the motion from toe to heel. The knife during the honing should be fitted in a knife-holder, and the back of the blade protected from the hone by a knife-back. The knife-blade must be kept free from grease and dirt, and nothing must be permitted to touch its edge. When many sections are cut at one sitting frequent stropping is necessary. When the cutting is finished the knife should be removed from the microtome, carefully cleaned and dried, and placed in its box. The slide and other bearings of the microtome should be well-oiled with the best microtome oil, and kept free from dust.

1. Cutting of Fixed Tissues Without Imbedding or Freezing. It occasionally becomes necessary in pathologic work to cut tissues directly upon the microtome without freezing or imbedding. This can be done satisfactorily in the case of very firm substances, such as amyloid liver and spleen, etc. The sectioning is done in the wet, using 80 per cent alcohol, as in celloidin cutting. A large celloidin knife should be used, and the blade should be flooded in alcohol. A large piece of the tissue, or, better, several pieces, are clamped in the object holder with only a small layer of tissue above the holder. The knife-blade is placed at the least possible angle to the pieces of tissue. To obtain sections of 15-20 microns in thickness the object-holder is raised each time about 25-30 microns. The sections will vary in thickness, some thin, others very thick.

2. Sectioning of Frozen Fixed Tissues. Tissues fixed in formol, Müller’s fluid or Orth’s fluid may be frozen directly on the freezing-microtome without previous washing. For other fixations previous washing is necessary. Tissues in alcohol must have the alcohol thoroughly washed out before freezing. Celloidin blocks may also be cut by freezing after the removal of alcohol. The fixed tissues cut to the proper size are placed on the object-holder of the freezing-microtome in a drop of saturated gum-arabic and the freezing carried out as directed above. (See Page [214].) As the freezing causes little or no damage to fixed tissue, the frozen sections may be transferred directly to alcohol, or floated out on the dilute molasses or sugar-dextrin solution, and stained separately or in sheets, according to the directions given in the next chapter.

3. Sectioning of Celloidin Blocks. The celloidin blocks are fastened in the object-holder of the microtome, and the knife-blade (a longer and broader one than for paraffin-sectioning is advisable) set nearly parallel with the longitudinal axis of the microtome, so that the cutting-edge, striking the block at a very slight angle, will be utilized for the greater part of its length in cutting entirely across the surface of the block. Both object and knife-blade must be kept constantly wet with 80 per cent alcohol; a broad camel’s-hair brush or a drip-bottle can be used for this purpose. The sections, as they are cut, are transferred from the knife to 80 per cent alcohol by sweeping the finger or brush from above toward the edge of the blade. Care must be taken not to strike the brush against the edge of the blade, and this can be avoided by using the brush always with a downward stroke. As the sections are cut they are guided onto the knife-blade by means of a fine-pointed camel’s-hair brush, if the sections show any tendency to curl. They should be smoothed out at once on the knife, and then transferred to the dish of 80 per cent alcohol. Sections 7-10µ thick are easily cut in celloidin, if the process of imbedding has been carefully carried out.

To prepare serial-sections from celloidin blocks the sections as they are cut must be arranged in their order on the knife-blade and thence transferred in this order to a slide or glass-plate, to which they are either fastened so that they cannot become loose during the staining process, or they are fastened together by a film of celloidin and stained in one piece. To fasten the celloidin sections to the slide Mallory and Wright suggest the cutting of the block in 95 per cent alcohol, and the arrangement of the sections in their order on the knife, whence they are drawn on to a dry, clean and numbered slide. Ether vapor from a half-full bottle of ether is then poured over the slide, slowly, to flatten out the sections and fasten down the frilled edges. The slides are then transferred to 80 per cent alcohol to harden the celloidin, and they are kept in this solution until ready for staining. Celloidin sections may also be fastened to the slide by albumin glycerin (equal parts white of egg and glycerin with a crystal of phenol or thymol). The celloidin sections, as they are cut, may be transferred to a dry gelatinized slide or plate (16 grms. gelatin in 300 cc. warm water), and are then covered with a thin celloidin; the slide is then placed in water at 50°C., the gelatin dissolves and the celloidin film floats off; the latter is then stained as one section. Serial sections of celloidin-blocks may also be arranged upon slides or plates covered with a coating of the Schmorl-Obregia sugar-dextrin solution or diluted New Orleans black molasses; the slide or plate is flooded with absolute alcohol; drained; a thin celloidin is then poured over it; it is immersed in warm water and the celloidin-sheet containing the sections is liberated, and is then stained as a whole or cut into strips as desired. The molasses method is the cheapest and simplest method, and much to be preferred to the Weigert closet-paper method, by which the sections are arranged upon a strip of moist closet-paper which is either held upon the slide or kept upon a piece of blotting-paper wet with 80 per cent alcohol. A slide or glass-plate is covered with a thin layer of celloidin and the strip of paper containing the sections is laid upon the celloidin surface, the sections down, so that these stick to the slide as the paper is carefully peeled off. The preparation is dried with filter-paper and a thin layer of celloidin poured over the slide and sections. The celloidin is then hardened in 80 per cent alcohol and the sections stained on the slide; or the celloidin film is removed by immersion in warm water, and then treated as one section. Celloidin films may be marked with a brush dipped in a water solution of methylene-blue. This should be done as soon as they are made. Langhans’ method is advised for the remounting of serial sections from tissues stained in bulk and imbedded in celloidin. The sections are cut in origanum-alcohol (1 part absolute alcohol to 3 parts origanum oil) and placed on a slide in a layer of origanum oil, blotted and mounted in balsam. Should sections become milky in the oil, renew the latter until they are cleared.

4. Sectioning of Paraffin Blocks. The paraffin block trimmed to the proper proportions, leaving a rim of paraffin about 2-3 mm. wide all about the tissue, is fastened to the wooden block by melting the under side of the paraffin by drawing a hot knife across it, and then immediately pressing the block upon the wood. The wooden block is then clamped into the object-holder. If desired the paraffin block can be fixed directly to the metal plate of the object-holder in the same manner. The block is raised until the level of the edge of the knife is reached. The knife is placed transversely or at a slight angle, and the cutting done with a relatively small portion of the edge. The paraffin block may be breathed upon to warm slightly the upper layer when a hard paraffin is used, and the knife is then drawn carefully through the block, guiding the section onto the knife-blade by means of a fine-pointed camel’s-hair brush held in the left hand. It is preferable, I think, to have a wider rim of paraffin at one corner of the block and to place the block with that corner at a slight angle to the knife, so that the edge of the blade will first engage the block at that point. The tip of the camel’s-hair brush moistened in water catches the corner of the section as the knife begins to cut, and pushes it over onto the blade, thus holding the section flat and preventing curling. When entirely cut through, the section is removed from the knife by the brush, still holding it at the corner first touched, or if necessary it is again moistened and applied to the upper side of the section, lifting the latter off the knife. The block is trimmed down to the proper level by cutting thick sections first, then sections of the desired thickness when the level of the entire block is reached. The sections are then transferred, with their shiny side down, just as they come off the knife, to a slide, sheet of paper, warm water, warm molasses or sugar-dextrin solution, or 70 per cent alcohol, and treated further according to the directions given in the next chapter. The presence of ridges on the cut surface of the block is an indication that the knife needs stropping. The knife must cut the paraffin, not scrape it. Crumbling of the paraffin is the result of imperfect infiltration during the imbedding process; water, alcohol, aniline oil or xylol may be left in the tissue, the paraffin may contain oil, or the cooling process may have been delayed. Curling of the sections can be prevented by using a sharp knife and slightly warming the surface of the block by breathing upon the block, use of a warm knife or spatula, heat-focus, etc. For ordinary work paraffin sections 5-7µ thick are easily obtained; for especial work 1-2µ sections may be obtained by especial care in imbedding, using graded paraffins and finally imbedding in a 56°C. paraffin. The pieces of tissue should be small, and the sections may be cut with a knife wet with water or alcohol. Serial paraffin sections are easily obtained; in fact, paraffin imbedding is by far the best method for serial cutting. To cut ribbons of sections upon the ordinary slide microtome the block must be clamped into the object-holder so that the edge of the block facing the knife, as well as the opposite side of the block, is parallel with the knife-edge. The knife should be placed at right angles to the microtome. If the paraffin has the right consistency the edges of the sections as they are cut will adhere, and a ribbon of serial sections will be pushed up over the knife. This ribbon can be cut into pieces of the desired length and mounted according to the directions given in the next chapter. The Minot automatic rotary microtome is especially well adapted for the cutting of ribbon sections. Failure of sections to adhere to each other is usually the result of too hard consistence of the paraffin, and this can be remedied by warming slightly, according to the method given above. If the paraffin is too soft the sections fold together. The block may be cooled in ice-water, or the sections may be cut with a cooled knife. Paraffin knives so constructed that they may be cooled by a stream of ice-water are supplied by dealers in microtomes. The conditions essential for successful paraffin cutting are perfect infiltration and imbedding, sharp clean knives and a certain degree of skill in manipulation that can only be secured by practice.

CHAPTER XXIV.
THE PREPARATION OF MOUNTED SECTIONS.

Sections of fresh material, unimbedded or imbedded tissues must be treated by a series of processes before they are finally permanently mounted and ready for use. These processes in general are: preparation for staining, staining, differentiation, washing, dehydration, clearing and mounting. The general procedure will be modified to some extent by the character of the tissue, manner of preparation (fixed or unfixed, imbedded or unimbedded, unstained or stained), character of stain (affected by alcohol, xylol, etc.), and the mounting agent (glycerin, balsam, damar, colophonium). Two or more of these steps may be combined in one; the same agent may differentiate, dehydrate and clear. Several stains may be combined in one solution, or it may be necessary to use them in succession. The very greatest variation is possible in pathologic technique; in fact, practically every laboratory worker modifies methods according to the light of his individual experience. The really important thing is to be master of the method, and not allow the method to control the situation. One of the greatest attractions about laboratory work is the infinite possibility of variation and improvement of methods and the invention of new ones.

I. PREPARATION FOR STAINING.

a. Frozen Sections of Fresh Tissue. Frozen sections of fresh tissues, as well as those obtained by the single or double razor, may be stained by floating the section on a slide, staining it directly (carbol-kresyl-echt-violett or carbol-thionin), examining in the stain or washing, dehydrating, clearing and mounting; or the section may be fixed to the slide with molasses or sugar-dextrin solution, covered with a celloidin-film, and treated according to the methods followed for paraffin or celloidin sections. Sections of fresh tissue may be fixed in formol or alcohol, and then treated by the same methods as celloidin sections. (See also Page [220].)

b. Sections of Unimbedded Tissues. These may be handled for staining in the same way as paraffin, celloidin or fresh-tissue sections, either when sectioned directly or after freezing. The sections may be stained directly, on the slide, cover-slip, or in the staining solution, or they may be transferred into celloidin sheets by the same methods employed in the preparation of paraffin sections.

c. Celloidin Sections. These may be transferred from water or alcohol directly to the stain. It is not necessary to remove the celloidin. If not stained soon after cutting they should be preserved in 95 per cent alcohol. Celloidin sections may be stained on the slide by simply blotting the section firmly on the slide, without permitting it to become dry, and manipulating it carefully through the various solutions; or the section may be fixed to the slide by the use of 95 per cent alcohol, ether-vapor and fixation in 80 per cent alcohol; or the section may be fixed to the slide by the methods given above under the cutting of serial sections of celloidin blocks. The most common method of preparation of celloidin sections for staining is to transfer the sections from alcohol into water to straighten them out, and then to transfer on the spatula into the stain. For the treatment of serial celloidin sections see above.

d. Paraffin Sections. Paraffin sections may be stained directly without removing the paraffin. This is especially advisable in the staining of tubercle-bacilli and in other cases where the use of alcohol is to be avoided. For many stains this method cannot be used. The sections as they are cut are floated directly into the warm stain, on which they flatten out, and are then transferred to the other solutions on the section-lifter, finally dried on the slide, in the incubator or over the flame, cleared in xylol and mounted in balsam. Paraffin sections may also be stained without removing the paraffin by being transferred directly from the knife on to 80 per cent alcohol, stained, washed, dehydrated in absolute alcohol or by drying, cleared and mounted. The section is transferred from one solution to another on the slide or spatula. The paraffin is removed during the clearing in xylol in both of these methods. The treatment with xylol must be on the slide, else the section may go to pieces. The staining of the section in the paraffin usually takes more time than staining after the paraffin has been removed, but the process can be hastened by heating the stain.

Slide and Cover-slip Preparations. Paraffin sections may be affixed to a slide smeared with a thin film of albumin-glycerin (equal parts of filtered beaten white of egg and glycerin, with crystal of phenol or thymol, or 1 grm. of sodium salicylate to 100 grms. of the mixture as a preservative). A drop of fixative is placed upon a clean slide, and is rubbed over the slide in a fine film with the back of the finger. The dry paraffin section with glossy side down is placed upon the smeared slide, flattened with a brush and then pressed firmly against the slide with the ball of the thumb. The albumin-fixative is then coagulated in the incubator or over the flame; the paraffin is melted over the flame without over-heating the section and the slide covered at once with xylol to remove the paraffin. It is then put into 95 per cent alcohol, thence into the stain; and after staining, the section is washed, dehydrated, cleared and mounted. Cover-glass preparations of paraffin sections are made by floating the sections with glossy side downward on warm water (just below the melting-point of the paraffin) until they straighten out and are perfectly flat. They are then floated on to cover-slips covered with a thin film of albumin-glycerin, the albumin having previously been coagulated by passing the smeared cover-slips through the flame quickly so that they do not scorch or burn. The cover-slips with the adherent sections are then placed in the incubator for 12 hours. The paraffin is then removed by xylol, the xylol is washed out in 95 per cent alcohol, and the cover-slips are then carried through the processes of staining, washing, dehydration, clearing and mounting. The cover-slips must be handled with forceps and the section side should always be uppermost. Slides covered with a film of albumin-glycerin may be used instead of cover-slips. The albumin-glycerin film may be omitted, and the sections with glossy side down floated in warm water on to clean covers or slides; the water is drained off and the slides or covers are put in the incubator for 12 hours. Sections adhere fairly well by this method (capillary attraction method). Bubbles are removed by careful heating. Serial ribbons of the size desired can be floated and mounted on slides by the albumin-glycerin or the capillary attraction method.

By far the best method of preparing paraffin sections for staining is the molasses plate method, a modification, originating in my laboratory, of the Schmorl-Obregia sugar-dextrin method. When many sections are to be stained at once it is the most convenient method and gives uniform results. In the preparation of sections for class-work it has no equal. It can be used also for giving out unstained sections. When many sections must be stained in diagnostic work the method saves much time and labor. Fifty sections can be stained as easily as one. It combines all the advantages of the celloidin and paraffin methods, as does the Schmorl-Obregia sugar-dextrin method, but is much cheaper than the latter.

Schmorl advised the use of a sugar-dextrin solution (cane sugar solution [1:1] 300 cc., 80 per cent alcohol 200 cc., yellow dextrin solution [1:1] 100 cc.) to be run over a perfectly clean glass plate or slide until the entire surface is covered with an even layer. The paraffin sections as they are cut are arranged in order on the wet plate, and when the plate is full, it is heated sufficiently to flatten and smooth the sections. The plate is then placed in an incubator for 3-12 hours to harden and dry. When dry it is immersed in xylol to take out the paraffin, then treated with absolute alcohol for 10-15 minutes, the alcohol drained off, and the plate covered with a thin layer of celloidin (celloidin or photoxylin 10, absolute alcohol 100, ether 100). As soon as the celloidin sets (1-2 minutes) the plate is immersed in warm water and the celloidin film containing the sections is detached. It can now be carried through the staining, washing, dehydrating and clearing solutions as one section, and in the clearing solution cut into strips or single sections, as desired, for mounting. Huber and Snow improved the method greatly by floating the paraffin sections directly on to warm dilute sugar-dextrin (a 10 per cent solution of Schmorl’s stock-solution will suffice), and plating the sections directly from the latter. This method of using the dilute solution is less expensive, much cleaner, and saves time in drying in the incubator. The results are in every way better than with the Schmorl solution in full strength. The formation of bubbles and crystals is almost wholly prevented, and less dust is caught on the plate. In my laboratory we have modified the method still further by using a 10 per cent solution of New Orleans black (or baking) molasses instead of the more expensive sugar-dextrin solution. As the molasses costs but 20 cents a gallon, a gallon of the dilute solution costing 2 cents can be used indefinitely if fermentation be prevented by a crystal of phenol or thymol. The paraffin sections are floated on to this dilute molasses solution warmed sufficiently to smooth out the sections; 4 × 5 glass plates (old negatives) thoroughly cleaned and kept in alcohol are immersed in the warm molasses solution and the sections arranged on them as desired, lifting out of the solution that part of the plate covered with sections as they are drawn upon it. As soon as the plate is covered it is drained, and is then flooded with absolute alcohol. After 1-2 minutes the alcohol is drained off and the plate flooded with thin celloidin, which is allowed to set for a minute or so, and the plate then immersed in warm water in which the celloidin film containing the paraffin sections is detached. This film is then handled by catching it at the two corners of one end with the fingers, or better still by a pair of forceps held in each hand. The film is put first into xylol to remove the paraffin, then into 95 per cent alcohol, then into water and thence into the staining solution. After staining the film is washed, dehydrated and cleared, and in the clearing solution is cut into strips or single sections by means of the wheel-shaped paper-cutter used by paper-hangers. The pieces are then mounted. A dilute sugar-dextrin solution can be used instead of the molasses-solution, but the latter is much cheaper and does just as well. Aside from this advantage our method of transferring the paraffin-sections into the celloidin film without first removing the paraffin saves a great deal of time, as it is not necessary to wait for the plates to dry in the incubator. The same method can be applied to the staining of single paraffin sections on the slide. The conversion of the paraffin section into a celloidin preparation without any loss of time for drying is so quickly and easily carried out that I advise it above all others. The same method may also be applied to the staining of fresh and fixed tissues cut on the freezing microtome or sectioned without imbedding. The success of the plate-method will depend largely upon the state of the glass-plates when put into the molasses solution. They must be perfectly clean or the celloidin sheet will not separate well. It is best to keep them in alcohol until they are needed. The celloidin must be of the right consistency, the layer must be thin, and cover the entire plate uniformly. It must not be allowed to harden too much before immersion in water or it will be tough and will shrink. Handling of the celloidin-sheets with the bare hands is not advisable because of the large number of epithelial cells adhering to the celloidin. The sheets are easily changed from one solution to another by catching them with forceps; the use of a glass-plate to transfer them is not necessary. When it is desired to preserve sections for future staining the celloidin sheet containing the paraffin-sections can be kept in 80 per cent alcohol indefinitely.

Note:—If the glass-plate is numbered with a blue wax-pencil after the paraffin sections are floated on, the marking will be transferred to the celloidin sheet, and the latter will retain the marking through all solutions.

II. STAINING AND DIFFERENTIATION.

Staining is necessary to bring out clearly the constituent elements of the tissues and their relations with each other, and for the demonstration of histologic structures or chemical substances that would otherwise be nearly or wholly invisible. The technique of staining depends upon the fact that stains or dyes possess certain affinities for the tissue-elements or for certain simple or complex substances present in the tissues (microchemic reactions). These affinities vary greatly with the dye. Some dyes have an affinity only for single constituents of the tissue (elective or specific stains); others have an especial affinity for the nucleus (nuclear stains), others stain all the tissue-constituents diffusely (diffuse or protoplasmic stains). There are but few pure elective stains for single tissue-elements; the majority of stains will stain more than one of the tissue elements, but may show an especial affinity for certain ones. As a result of these variations in the affinities of dyes for the constituents of the tissues it becomes possible to manipulate the dyes or to combine them in such a way that a specific differentiation of many tissue-elements is possible through the use of different methods of staining. These methods are based in part upon the use of different mordants, the employment of several stains in combination or in succession, the mixture of stains to form a new staining compound, the phenomenon of metachromasia, the differentiation of certain tissue-elements by the removal of the stain from the structures for which it possesses a weaker affinity, and by the employment of different microchemic reactions. The two most commonly employed methods are the progressive, in which the stain is allowed to act until the affinities of certain tissue-elements have been satisfied when the staining process is interrupted; and the regressive, in which the tissue is over-stained, and the dye withdrawn from the tissue-elements for which it possesses the weakest affinities leaving the other elements stained. This latter process is usually called “differentiation,” and the chief substances used for such differentiating are dilute acid, acid alcohol, acid stains, aniline oil, aniline-xylol and alcohol. Some workers use the regressive method for such simple stains as hæmatoxylin, overstaining, and then differentiating with acid alcohol before counterstaining with eosin. The results obtained in this way are much less satisfactory than is possible with the progressive method.

Tissues may be stained in the body during life (intravital staining), or immediately after removal from the body (supravital or survival staining), either before or after sectioning. (See Page [217].) Fixed tissues may be stained in bulk or in sections.

Staining Tissues in Bulk.

This method is not often used in pathologic work. The fixed and hardened tissue is cut into small pieces, placed in the staining solution for several days, washed thoroughly, dehydrated in alcohol, imbedded, cut, and mounted without further staining. Alcoholic solutions penetrate best; hæmatoxylin, hæmalum, carmine and alcoholic solutions of the aniline stains may be used. Metallic impregnation (gold or silver salts) of fresh or fixed tissues is but little used in pathology. (See Staining of Nervous System, and Spirochætes.)

Staining of Sections.

Celloidin sections are lifted from water or alcohol into the stain by the needle or section-lifter. The use of the latter is advised, as by it the section can be floated flat on to the staining solution. When many celloidin sections are to be stained at once they can be stained in small tea-strainers and transferred in these from one solution to another. Paraffin sections may be floated directly on to the stain without removing the paraffin; or they may be stained on the slide or cover-glass after removing the paraffin, the stain being dropped on to the section, or the slide or cover-slip is immersed in the stain. Special staining-dishes for the staining of paraffin sections on slides and covers can be obtained. Paraffin sections transferred to celloidin sheets by the plate method can be put into the staining-solution while on the glass-plate, or the films can be detached and transferred from one solution to another by means of forceps. This is the easier way, and it is not necessary to touch the films with the fingers.

General Rules for Staining.

1. The stain should be filtered just before being used, in order to remove precipitates, moulds, etc. Unless they have been diluted most of them can be used over and over again, hence after using they should be filtered back into the stock bottle.

2. A liberal amount of stain should be used. Slides and cover-slips are given enough stain to cover completely the section, when the staining is done on the slide, or they may be immersed in staining-dishes. Plates and celloidin sheets should be stained in large trays. Sections and celloidin films should be flat without folds or wrinkles, and they should not touch one another when several are stained at the same time. Transference of the section from water or dilute alcoholic solutions to dilute or stronger alcohol respectively for a moment and then back again will usually straighten out curled or wrinkled celloidin sections.

3. Stain until the section is properly stained. Control this by removing it from the staining-solution and examining it in water on a glass-slide without a cover-slip, using the low-power. Sections will always appear more deeply-stained when cleared than when examined in water, hence due allowance should be made. Celloidin sheets can be examined on glass-plates. The time-limits given in staining methods are only approximate; no absolute rules can be laid down as to the length of time necessary to obtain a good stain. The methods of fixation and hardening, age of the tissue, age of the stain, etc., affect the staining power. Some stains lose their staining-power after a time; others require a period of ripening before they yield the best results. As a rule staining may be intensified or hastened by staining in the incubator or at a higher temperature, by concentrating the stain, or by the use of such substances as aniline oil. When differentiation is necessary the process should also be controlled by frequent examination of the section, as above for staining. Usually the section can be examined in the differentiating fluid.

III. WASHING.

Thorough washing after staining is necessary after nearly all stains. The washing should usually be done by soaking the sections in several changes of distilled water, although tap-water, alcohol, alum-water, and other solutions may be used to intensify the staining effects. When this is done a final washing in distilled water or alcohol is usually necessary. Differentiating fluids should always be thoroughly removed from the section before mounting. Sections should not be allowed to lie in wash-water that is colored by the stain; as soon as the wash-water becomes colored it should be replaced by fresh. When sections are left lying in the wash-water for some time the vessel containing them should be covered to prevent the settling of dust on the sections, as it is practically impossible to remove from the latter dust or other precipitates that may become attached to them. Some stains give better results after long washing; others are easily washed out if the sections are left standing in the wash-water. The time-limits of washing will depend upon the character of the stain employed.

IV. DEHYDRATION.

Dehydration of the sections is usually produced by passing them through two alcohols, 80 per cent and absolute, or 80 per cent and 95 per cent. For certain clearing reagents (xylol) it is necessary to use absolute alcohol. When carbol-xylol is used as a clearing reagent absolute alcohol is not necessary, and 95 per cent can be used instead for the second dehydrating solution. Usually a minute in each alcohol is sufficient for the dehydration of single sections. If dehydration with alcohol is not desirable because of its action on the stain it is possible to dehydrate and clear in xylol by repeatedly blotting the section with absorbent paper, covering the section several times with xylol and then blotting. The section should never be allowed to become perfectly dry. Dehydration with alcohol may also be avoided by staining paraffin sections without removing the paraffin, drying in the incubator or over the flame, removing the paraffin in xylol, and mounting. Imperfect dehydration is shown by the presence of white spots or a milky cloud in the section when it is put into the clearing fluid.

V. CLEARING.

After dehydration, sections must be cleared in some solvent of balsam before they can be mounted in the latter medium. When 95 per cent alcohol has been used for the final dehydration the sections may be completely dehydrated and cleared at the same time by the use of carbol-xylol (xylol, 3 parts; melted crystals of carbolic acid, 1 part; add melted carbolic acid to the xylol to prevent formation of crystals). The sections (on the slide or cover-slips, in celloidin sections or films) are transferred from the alcohol, draining or blotting off excess of the latter, into the carbol-xylol, and left until perfectly clear. This can be most easily determined by viewing the sections against a dark background. Carbol-xylol cannot be used for sections treated with aniline stains. These are dehydrated in 95 per cent alcohol, and the final dehydration and clearing accomplished by repeatedly placing xylol upon the slide and blotting it out until the sections are transparent. Turpentine, chloroform, benzine, toluol, the oils of bergamot, cloves, thyme, lavender, origanum cretici, and cedarwood, aniline oil, and various mixtures of these oils are also used as clearing agents. The majority of these will clear from 95 per cent alcohol, but not so readily as carbol-xylol; they have individual disadvantages of taking out the eosin, affecting aniline colors, dissolving celloidin, making sections brittle, slow action, clinging odor, etc. Chloroform and benzine may be used for clearing osmic-acid preparations; oil of turpentine is also good for clearing sections stained with kresyl-echt-violett, and Wright’s blood-slain. With but few exceptions carbol-xylol and xylol meet all requirements better than any other clearing reagents. There is but one disadvantage in the case of carbol-xylol; some of the phenols in the market cause a fading of eosin and hæmatoxylin stains. DeWitt has shown that this fault can be corrected by redistillation, stopping the distillation as soon as the temperature begins to rise above the constant boiling point of the phenol; or the carbol-xylol that fades the stains can also be corrected by supersaturating it with a mixture of sodium bicarbonate one part, and sodium-potassium tartrate two parts. Sections kept in xylol or carbol-xylol should be protected from dust and evaporation; it is not a good plan to keep them in these solutions for more than 24 hours.

VI. MOUNTING.

Permanent mounts are made in glycerin, potassium acetate, lævulose, glycerin gelatin, balsam, damar or colophonium. For celloidin and paraffin sections a solution of Canada balsam in xylol is most commonly used for mounting. Celloidin sections (celloidin films are best cut into strips and single sections when in the carbol-xylol; the wheel-shaped paper-cutter used by paper-hangers is the best instrument for this purpose) are lifted onto the slide from the clearing-fluid; folds or wrinkles in the celloidin are straightened or removed by cutting the celloidin at right angles to the section in order to relieve the tension, and the section is then blotted firmly against the slide by means of a pad of absorbent paper. The greatest care should be taken to prevent wrinkling, folding or turning over of the edge of the section. As soon as the pad is removed a drop of balsam is placed upon the section and the cover-glass put over it. There should be just enough balsam used to fill the space between cover-slip and slide, so that air-bubbles are not formed. The balsam must not be so thin that the cover-glass will float about on the liquid, or so thick that it does not spread well. In the latter case warming the slide may cause it to spread more readily, but care must be taken not to injure the stain by over-heating. Paraffin sections on the slide are similarly blotted and covered with balsam and cover-glass; those on cover-slips are blotted between folds of absorbent paper and immediately placed with section-side downward upon a drop of balsam that has been put upon the slide.

Xylol-damar may be used in place of Canada balsam; it is cheaper and colorless, but it tends to become cloudy. Colophonium is the cheapest of the three and has but little color; it is highly recommended by many workers. In a xylol-solution it may be used for aniline stains; a chloroform solution is advisable for the mounting of osmic-acid preparations; while a solution in turpentine and shellac is recommended for Weigert’s neuroglia method, Wright’s blood-stain, and other special staining methods.

Glycerin, potassium acetate, laevulose and glycerin-gelatin are used for the preservation of amyloid-, mucin- and fat-stains, as well as for other preparations that do not permit the use of alcohol and xylol. Glycerin-gelatin is probably the best medium for this purpose. It is made according to Kaiser by soaking 7 grms. of gelatin for 2 hours in 42 cc. distilled water, then adding 50 grms. glycerin and 1 grm. carbolic acid; the mixture is warmed 10-15 minutes, stirring constantly, and filtered while hot. It is also made by taking water, 200 cc.; gelatin, 20 grms.; powdered white shellac, 2 grms.; Farrant’s solution (gum arabic, glycerin, solutio acidi arsenicosi conc., aa. 30.0 grms.), dissolve by warming, and filter while warm. To mount in this medium the section is placed on the slide, and blotted with absorbent paper. A drop of warm glycerin-gelatin is then placed on it and the cover-slip affixed. The drop spreads evenly beneath the cover-glass and becomes solid as it cools. Mounts in glycerin, glycerin-gelatin, potassium acetate and laevulose must be cemented around the borders of the cover-slip with asphalt, wax, paraffin, gold size, etc., using a brush or glass rod for this purpose.

VII. SUMMARY OF METHODS OF PREPARATION OF CELLOIDIN SECTIONS.

1. Fixation of the tissues in alcohol, formol, etc.
2. Wash 24 hours, when necessary.
3. After-harden in 80 and 95 per cent alcohols for 1 to several days.
4. Complete dehydration in absolute alcohol for 24 hours.
5. Equal parts of pure ether and absolute alcohol, 24 hours.
6. Thin celloidin, 1-3 days.
7. Thick celloidin, 1-3 days.
8. Block. Harden block in 80 per cent alcohol.
9. Cut; keep sections in 80 per cent alcohol.
10. Stain; differentiate.
11. Wash thoroughly.
12. Dehydrate in 80 and 95 per cent alcohols.
13. Final dehydration and clearing in carbol-xylol.
14. Place on slide and blot with absorbent paper.
15. Mount in xylol-balsam or xylol-colophonium.

VIII. SUMMARY OF METHODS OF PREPARATION OF PARAFFIN SECTIONS.

1. Fixation of the tissue in alcohol, formol, etc.
2. Wash 24 hours, when necessary.
3. After-harden in 80 and 95 per cent alcohol for 1 to several days.
4. Complete the dehydration in absolute alcohol for 24 hours.
5. Aniline-oil until tissue becomes transparent.
6. 1st. Xylol, ½ hour, to remove aniline oil.
7. 2nd. Xylol, 1-2 hours, until translucent.
8. 1st. Paraffin (52°C.), ½ hour in oven, to remove xylol.
9. 2nd. Paraffin (52°C.), 1-12 hours in oven, until infiltrated.
10. Imbed and block; cool quickly.
11. Cut sections; mount on slides or covers.
12. Remove paraffin in xylol.
13. Remove xylol in absolute alcohol.
14. 80 per cent alcohol, for a few minutes.
15. Stain; differentiate; wash.
16. Dehydrate in 80 and 95 per cent alcohols.
17. Clear in carbol-xylol.
18. Mount in Canada-balsam or colophonium.

IX. ARTEFACTS IN MOUNTED SECTIONS.

A mounted section, after passing through the various stages indicated above, must of necessity present some appearances that are the result of the technical methods employed. The number and degree of such artefacts will depend upon the character of the methods employed and the care exercised in their performance. The trained observer ignores the presence of artefacts as having nothing at all to do with the significance of the section itself; but to the beginner in microscopic work they often appear to be the most important thing in the preparation, and are given a pathologic interpretation. How frequently do we see students, undergraduates and postgraduates, take up a section and pick out a fold, wrinkle, tear, staining-defect, precipitate, dirt, etc., as pathologic features! It is necessary, therefore, for the student to acquaint himself with the nature of artefacts so that he may ignore them and not give them an incorrect interpretation. The most important artefacts are as follows:—

1. Artefacts due to fixation (mercuric, chromic, osmic, etc., precipitates; alterations in blood-pigment due to formol; loosening of cells from basement membrane due to contraction [kidney tubules, etc.: loosened endothelium in blood-vessels, particularly confusing to students]; destruction of red blood cells, as in alcohol fixation; poor staining due to over-fixation).

2. Artefacts due to hardening (contraction, desquamation of cells, etc.).

3. Artefacts due to imbedding and cutting (tears, holes, ragged edges, irregular thickness, knife-streaks, compression of soft structures, dislocation and tearing-out of firm tissues or material, wrinkles, folds, etc.).

4. Artefacts due to poor staining (uneven, spotted or streaked staining, overstaining, understaining, poor differentiation, precipitates, insufficient washing, fading, poor contrasts, defective staining due to presence of paraffin, dextrin, etc., in section).

5. Artefacts due to poor mounting (imperfect dehydration and clearing, cloudiness, milkiness or opacity of section; folds; wrinkles; turned-over edges; tears in section caused by striking it with balsam-dropper or needle; air-bubbles; lack of balsam).

6. Dirt and foreign-material (opaque and translucent dirt, above or below section; coloring-matter in balsam; ink; pigment from pencil; cotton-, silk, wool-, linen-, vegetable and paper-fibres, hairs, desquamated squamous epithelium, portions of insects, etc.)

CHAPTER XXV.
STAINS AND STAINING METHODS.—NUCLEAR AND PROTOPLASMIC STAINS.

THEORIES OF STAINING. The exact nature of the process of staining has not yet been determined. Various theories have been advanced, explaining the affinity of the tissues for certain dyes, on the ground of a chemical, mechanical or chemicophysical action. The chemical theory assumes the formation of an insoluble compound through the chemical combination of tissue and stain; the physical theory is based upon the assumption that the process is purely physical or mechanical, while the chemicophysical theory holds that it is neither purely physical nor purely chemical. The process is not controlled by the molecular weights alone of the substances concerned, but does depend upon the conditions controlling the formation of solutions in general. Therefore, the theory most widely accepted at the present time is the solution-theory, which assumes that the staining-process is a solution of the dye in the tissue, and that the stained tissue-element is a fixed solution of the stain in its substance. This solution of the dye in the tissues may be a direct action between the two (direct or substantive stains); or it may be brought about only by the interaction of a third substance (indirect or adjective stains). The third substance is called a mordant, and the combination of the dye with the mordant is known as a lake. The mordant may be added to the stain or to the tissue, either before or at the time of staining. Many of the fixing-fluids are mordants, particularly those containing chromic acid or its salts. Alum, iron, and many of the metals are the most commonly used mordants. In a general way acid mordants are used for basic colors, and basic mordants with acid colors.

The stains most commonly used in pathologic work are:—

1. Natural Dyes:—Hæmatoxylin and Carmine.

2. Aniline Dyes:—a, Acid.—Eosin, erythrosin, acid fuchsin, orange G, picric acid, sudan III, and scarlet R (Fett-ponceau).

b, Basic.—Methylene blue, methylene violet, thionin, toluidin blue, kresyl-echt-violett, methyl violet, gentian violet, crystal violet, basic fuchsin, dahlia, aniline blue, methyl green, iodine green, safranin, Bismarck brown, and vesuvin.

In a general way it may be said that basic stains are nuclear stains, and acid stains are protoplasmic. Neutral stains are usually diffuse stains; when formed by the combination of acid and basic dyes they usually act as selective stains for some especial tissue-element or cell-constituent. Metachromasia, in its narrowest sense, is the term applied to that staining-phenomenon, in which a single-chemical entity gives different colors to different tissue-elements. In this sense iodine, in its action upon glycogen and amyloid, is a true metachromatic substance. The majority of the so-called metachromatic stains, however, do not possess true metachromasia, since their metachromatic powers are dependent upon a mixture of two or more dye-stuffs in the one compound. The most important stains of this class are gentian violet, methyl violet, crystal violet, dahlia, thionin, toluidin-blue, polychrome methylene blue, methylene azure, and kresyl-echt-violett. The chief chromotropic substances are amyloid, mucin, mast-cell granules and cartilage. Metachromatic reactions are at their best usually when examined in water; they are affected by alcohol and usually destroyed by carbol-xylol. Sections stained by metachromatic stains should be quickly dehydrated by absolute alcohol and blotting, and cleared in xylol, when mounted in balsam.

I. NUCLEAR STAINS.

1. Haematoxylin (C16H14O6) is an ether extract of the wood of Hæmatoxylon campechianum, a tree found in the West Indies and Central America. In itself not a dye, it becomes one of the most valuable when oxidized to hæmatein (C16H12O6), and combined with alum, iron or other mordants to form a lake. It then stains nuclei a deep violet-blue or black color that is practically permanent. Mucin, lime-salts, bacteria, and colonies of actinomyces are also stained varying shades of blue. If the staining process is prolonged the entire tissue, as well as celloidin, becomes more or less heavily stained blue. A pure nuclear stain is obtained by interrupting the stain at the right time (examine in water); or if the sections are over-stained they may be differentiated in acid alcohol (1 per cent hydrochloric acid in 70 per cent alcohol). Hæmatoxylin stains well after all fixing-solutions except osmic acid; some of its staining-formulæ stain slowly after fixation in Zenker’s fluid. On the whole, it is by far the best general nuclear stain for laboratory and diagnostic work. It is employed in numerous staining formulæ, the most useful of which are given here. These formulæ differ chiefly in the time of staining, “ripening” of the stain (oxidation of hæmatein), intensity of stain, necessity of differentiation, etc.

a. Böhmer’s Alum-haematoxylin.

Dissolve 5 grms. of hæmatoxylin crystals in 50 cc. of absolute alcohol; add this drop by drop, while stirring, to 1,000 cc. of a 1 per cent solution of potassium alum. Expose in open vessel to air and light for 1-2 weeks. Filter before using.

b. Hansen’s Haematoxylin.

To 200 cc. of alum-hæmatoxylin solution brought to the boiling-point add 2 cc. of a concentrated solution of potassium permanganate. Cool quickly; filter when cold. Can be used at once without further ripening. It tends to stain diffusely.

c. Delafield’s Haematoxylin.

To 400 cc. of a saturated solution of ammonia alum add a solution of 4 grms. of hæmatoxylin in 25 cc. of absolute alcohol. Expose mixture to air and light for 3-4 days; filter; then add 100 cc. of glycerin and 100 cc. of 95 per cent alcohol, and filter. Expose to light until solution is dark enough, then keep in tightly-stoppered bottle. It is a strong stain, and may be diluted with distilled water when desired. The solution keeps well.

d. Ehrlich’s Acid-haematoxylin.

Dissolve 2 grms. of hæmatoxylin in 100 cc. of absolute alcohol. Add this to a saturated solution of potassium alum in water 100 cc., glycerin 100 cc., and glacial acetic acid 10 cc. Allow mixture to stand for a week exposed to air and light; then filter. Keep in well-stoppered brown bottles. The solution stains best when it is six months old, and may be kept for several years. It does not overstain, and on the whole is more useful than Böhmer’s, Hansen’s or Delafield’s.

e. Mayer’s Haemalum.

Dissolve 1 grm. of hæmatein in 50 cc. of 90 per cent alcohol and warm. Add this solution to a solution of 50 grms. of potassium alum in 1,000 cc. of distilled water dissolved by heating. Mix warm, cool, and filter. With hæmatein no ripening is required, and the solution can be used at once. Hæmalum is a more precise nuclear stain, but stains more slowly than the formulæ given above.

It may be prepared directly from hæmatoxylin crystals by dissolving 1 grm. of hæmatoxylin in boiling water; add water up to 1 litre, and cool. Add 0.2 grm. sodium iodate and 50 grms. potassium alum, dissolving at room temperature; 50 grms. chloral hydrate and 1 grm. citric acid may be added to make solution keep better.

f. Mayer’s Acid-haemalum.

Add 2 cc. glacial acetic acid to 100 cc. of hæmalum solution. It stains the nuclei more precisely than hæmalum.

g. Weigert’s Iron-haematoxylin.

Dissolve 1 grm. of hæmatoxylin in 100 cc. of 96 per cent alcohol. Allow to ripen several days, but this solution should not be kept longer than six months. Make a second solution of 4 cc. of liq. ferri. sesquichlor. (German Pharmak. IV, sp. gr. 1,124), 1 cc. of concentrated hydrochloric acid and 100 cc. of water. Mix equal parts of each solution just before staining. The mixture stains well for 5-8 days, so that a quantity of stain sufficient for this time only should be made up. The two stock solutions are easily made and keep well. The nuclei stain quickly and deeply, and differentiation and long washing are unnecessary when hydrochloric acid is included in the second solution, as given above. After-staining with eosin, picric acid or the Van Gieson’s mixture gives better results with Wiegert’s iron-hæmatoxylin than with any other hæmatoxylin.

Method of Staining with Haematoxylin.

1. Stain 1-15 minutes, controlling progress of stain by examination of section in water, on slide, using low-power.

2. If sections are over-stained, differentiate in ½-1 per cent potassium-alum or in acid alcohol.

3. Wash thoroughly in tap water, until a good blue is obtained. Exposure to ammonia vapor or washing for a few seconds in lithium-carbonate solution will hasten the development of the blue color. If these reagents are used the section should afterwards be thoroughly washed in water.

(Stain with a plasma stain, if contrast is desired.)

4. Dehydrate in 80 and 95 per cent alcohols.

5. Clear in carbol-xylol.

6. Mount in balsam.

Over-ripened hæmatoxylins may stain reddish or even brownish, and too diffusely. In such cases the celloidin will be deeply stained. The addition of alum-water to the stain may counteract the fault. Alum-hæmatoxylins must always be filtered before using, as precipitates are constantly formed as the result of oxidation.

2. Carmine. Carmine is the coloring matter of cochineal, the dried bodies of the female coccus cacti, and is obtained chiefly from Honduras. The coloring principle is carminic acid (C2H22O12). When combined with alum, borax, lithium, etc., carmine gives a good, permanent nuclear stain, varying from reddish violet to deep scarlet. It is used chiefly in pathology to give a contrasting nuclear stain to the various pigments, and when specific blue stains have been used for fibrin, mucin, bacteria, elastic tissue, etc., or when a blue injection-mass has been used. Alum-carmine is the most precise nuclear stain. Differentiation with acid-alcohol is necessary after staining with borax- or lithium-carmine. Lithium-carmine is on the whole the best of the three for use as a contrast-color to the various pigments.

a. Alum Carmine.

Carmine ½-1 grm., 1-5 per cent alum solution 100 cc.; boil 20 minutes; cool; filter. Add crystal of thymol as preservative.

1. Stain 15 minutes to several hours.

2. Wash thoroughly in distilled water.

3. Dehydrate in 80 and 95 per cent alcohols; clear in carbol-xylol: mount in balsam. Nuclei are a light reddish violet; the plasma is slightly stained (muscle) or not at all.

b. Lithium Carmine.

2-5 grms. of carmine to 100 cc. of a cold saturated water solution of lithium carbonate; filter.

1. Stain 1-3 minutes; transfer directly to acid alcohol (1 cc. of hydrochloric acid to 100 cc. of 70 per cent alcohol) without putting section into water; differentiate ¼-6 hours, until nuclei alone retain the color. Control differentiation by examination on the slide in acid alcohol.

2. Wash thoroughly in water.

(Use plasma stain, as picric acid, if desired.)

3. Deyhdrate in 80 and 95 per cent alcohols; clear in carbol-xylol; mount in balsam.

c. Borax Carmine.

Dissolve by boiling 0.5-0.75 grm. carmine and 1-2 grms. of borax in 100 cc. of water. To the hot solution add 5 cc. of a O.5 per cent acetic acid until solution is deep red. After 24 hours filter and add crystal of thymol.

Stain and differentiate as with lithium-carmine, but leave the sections somewhat longer in the stain (15 minutes).

3. Basic Aniline Stains. The basic aniline stains are used as general stains for bacteria in sections, and at the same time stain the nuclei. Methylene blue and fuchsin are employed especially for this purpose. The metachromatic dyes (thionin, kresyl-echt-violett, etc.) are also used as nuclear stains in combination with their metachromatic reactions with mucin, amyloid, mast-cells, etc. Methylene blue is used in the study of the blood-forming organs, cell-inclusions, parasites in the tissues, etc. Safranin and fuchsin are used for staining mitotic figures. (See Staining of Mitoses.) Bismarck brown is employed sometimes in preparing sections for microphotography. Methyl green is an intense chromatin stain and is used in various combinations. Since these dyes are not very permanent, and are easily washed out in dehydrating and clearing fluids, as well as “running” in balsam mounts, they are rarely employed as pure nuclear stains in pathologic work.

When the basic-aniline stains are used a saturated water or concentrated alcoholic solution (1½-2 per cent in 40 per cent alcohol) may be employed as stock-solution and diluted 1:5 or 1:10, as desired. The sections are stained 3-5 minutes, then differentiated in absolute alcohol, cleared in xylol, and mounted in balsam. Methylene-blue is used by some workers as a nuclear stain contrasted with eosin for tissues fixed in Zenker’s solution, giving better effects with this fixation than the hæmatoxylins. Unna’s alkaline methylene-blue formula is employed (methylene-blue 1 grm., carbonate of potassium 1 grm., water 100 cc.). Dilute 1:10 or 1:5 for staining. Stain 10-15 minutes, wash quickly in water, differentiate in 95 per cent alcohol; dehydrate in absolute, clear in xylol, mount in balsam. When celloidin sections are used, 95 per cent alcohol may be used for dehydrating, blotting on the slide with several changes of xylol.

The nucleolus has an affinity for the acid stains. With the modifications of the Romanowsky methylene-blue-eosin methods the nucleolus stains red, the nucleus blue.

II. DIFFUSE OR PLASMA STAINS.

The most commonly used diffuse stains are eosin, erythrosin, acid fuchsin, orange G, and picric acid. They are practically never used alone, but are employed as contrast-stains to the nuclear stains. Eosin, orange G, acid fuchsin and picric acid may be used as counterstains for hæmatoxylin; picric acid is used as the best contrast to the carmines, while eosin and orange G are employed as counterstains for methylene blue. The combination of hæmatoxylin and eosin is by far the best general staining method for laboratory and diagnostic work, except for tissues fixed in Zenker’s solution; for these the combination of methylene blue and eosin is preferable. Ammonia carmine is sometimes used as a diffuse stain for bone and the central nervous system. The majority of the diffuse stains wash out easily in water, alcohol and xylol, hence sections thus stained should not be allowed to remain too long in these fluids.

a. Eosin. Two forms of eosin are obtainable, one soluble in water, the other in alcohol. Saturated solutions of both kinds should be kept as stock solutions and diluted as occasion demands. For use after hæmatoxylin a ½ per cent solution is advisable; with Zenker’s fixation a more dilute solution may be used, as tissues so fixed stain intensely in eosin. If used as a contrast-stain to methylene-blue, eosin is used first in a 5 or 10 per cent solution, as the basic nuclear stain takes out some of it. Some workers express a preference for the aqueous solution of eosin, others for the alcoholic; the alcoholic solution stains more uniformly and with less differentiation than the other. Eosin is particularly good as a contrast-stain for tissues containing red blood cells when fixed in formol, mercuric chloride or Zenker’s.

b. Orange G. Used in a 1 per cent water solution. Requires longer time for staining than eosin.

c. Acid Fuchsin. A saturated water solution is kept in stock and diluted as needed. Must be used in weaker solutions than eosin, as it more quickly overstains, and cannot be washed out so well.

d. Picric Acid. Keep in stock either a saturated water or saturated alcoholic solution, and dilute as needed. As it washes out more readily than eosin, the staining solution should be stronger than for the latter, and the sections somewhat over-stained to allow for some loss of stain. Picric acid gives a brownish tint to nuclei stained with hæmatoxylin or carmine, and will take out the stains completely, if allowed to act too long.

e. Ammonia Carmine. One grm. of carmine is dissolved, without heating, in 50 cc. of distilled water and 5 cc. of strong ammonia water. The fluid is then exposed in an open dish until the odor of ammonia is lost; it is then filtered. When ready for use dilute by filtering 1-2 drops into 20 cc. of distilled water.

III. COMBINED NUCLEAR AND DIFFUSE STAINS.

The diffuse or plasma stains may be combined with the nuclear in one staining solution, or used in succession. The latter method gives better results. The nuclei are stained first, the diffuse stain being used after the washing-out following the use of the nuclear stain, except in the methylene-blue and eosin method in which to obtain the best results it is necessary to stain with the eosin first. Nuclear hæmatoxylin may also be followed by a combination of plasma stains, as in the Van Gieson’s mixture of picric acid and acid fuchsin, Delépine’s mixture of rubin and orange, White’s erythrosin and picric acid mixture, etc. In these mixtures the different affinities of the plasma-stains give rise to differential or selective staining effects.

a. Haematoxylin and Eosin.

1. Stain in any one of the hæmatoxylins.

2. Wash thoroughly.

3. Stain in dilute water or alcoholic eosin until section is bright rose-red.

4. Differentiate eosin-staining, as desired, by rapid or slow washing in water.

5. Dehydrate quickly in 80 and 95 per cent alcohols.

6. Clear quickly in carbol-xylol. (If carbol-xylol takes out eosin too rapidly add some of the dry eosin stain to it. Hæmatoxylin-stained sections can be placed in such eosin-carbol-xylol and will take up the eosin beautifully.)

7. Mount in balsam.

Hæmatoxylin and eosin can also be combined in one stain, but the results are not as good as those obtained by successive staining.

b. Haematoxylin and Picric Acid.

1. Stain with Weigert’s iron-hæmatoxylin, or overstain with any other hæmatoxylin.

2. Wash thoroughly.

3. Stain in saturated water solution of picric acid, diluted one-half, until sections are a bright yellow. If left too long in the stain, the hæmatoxylin will become brown or may be wholly lost.

4. Wash, dehydrate and clear quickly, as for eosin. (Dry picric acid may be added to the carbol-xylol.)

5. Mount in balsam.

c. Haematoxylin and Acid Fuchsin.

Stain with hæmatoxylin, and after washing use a 1 per cent water solution of acid fuchsin, until section is sufficiently red; wash; dehydrate; clear; mount.

d. Haematoxylin and Orange G.

Stain with hæmatoxylin, and after washing use a 1 per cent water solution of orange G, staining ¼-3 hours. Treat otherwise as for eosin-staining.

e. Carmine and Picric Acid.

Stain with borax- or lithium-carmine; differentiate in acid alcohol and wash thoroughly. Then counterstain with picric acid, as for hæmatoxylin and picric acid. Carmine and picric acid may also be combined in one stain, as picro-carmine, but this is rarely used at the present.

f. Eosin and Methylene-blue.

(For tissues fixed in mercuric chloride or Zenker’s.)

1. Stain in a 5-10 per cent aqueous eosin for 20 minutes or longer, until a deep eosin-stain is obtained.

2. Wash out excess of eosin in water.

3. Stain in Unna’s alkaline methylene-blue, diluted 1-4 or 5 with water, 10-15 minutes.

4. Wash in water.

5. Differentiate in 95 per cent alcohol, keeping the section in constant motion to obtain a uniform decolorization. Control process under microscope. Wolbach advises the use of a 0.75-1.5 per cent solution of colophonium in methyl alcohol as a differentiating medium instead of 95 per cent alcohol. For tissues fixed in formol or alcohol a 10 per cent solution should be used.

6. When the background is pink, dehydrate quickly with absolute alcohol, or in 95 per cent by blotting on slide with xylol, until clear.

7. Clear in xylol.

8. Mount in balsam.

g. Van Gieson’s Method.

1. Stain in Weigert’s iron-hæmatoxylin, or overstain if any other hæmatoxylin is used. (Weigert’s gives the best results, as it does not decolorize so readily.)

2. Wash thoroughly.

3. Stain in Van Gieson’s mixture (acid fuchsin 1.5 grms., saturated water solution of picric acid [0.6 per cent] 150 cc. This mixture keeps well. Add 1 cc. of this stock solution to 10 cc. of saturated water solution of picric acid. Stain in this for 10 seconds).

4. Wash quickly; dehydrate in alcohol; clear in xylol or carbol-xylol; mount in balsam.

I have obtained the best results by making the Van Gieson’s mixture by taking an ordinary small staining-dish nearly full of saturated water solution of picric acid, and adding to this, drop by drop, sufficient saturated water solution of acid fuchsin to make the solution just dark enough so that the finger cannot be seen through the staining-dish. The hæmatoxylin-stained section is put into this mixture for a few seconds, until it appears to become lighter. The section is then washed in 95 per cent alcohol, dehydrated, cleared and mounted.

The Van Gieson method is extremely valuable in pathologic work, because of its varied differential reactions. The nuclei are brown or black, protoplasm is ochre-yellow, connective-tissue light red, voluntary and involuntary muscle yellow, axis-cylinders red, connective-tissue hyalin deep rose-red, epithelial hyalin yellow, orange or brownish, amyloid yellow or brownish pink, mucin yellow or brownish, fibrin yellow or brown, necrotic areas yellow or brownish, lime-salts brown to brownish blue or violet.

Combinations of rubin and orange (Delépine) and erythrosin and picric acid (Powell White) are also advised as differential combination stains, but are not so useful as Van Gieson’s. Other combination methods are to be found in the various modifications of the Ehrlich triple stain.

CHAPTER XXVI.
SPECIAL STAINING METHODS FOR DEMONSTRATION OF PATHOLOGIC CONDITIONS IN CELLS OR TISSUES.

I. AMYLOID. The best selective staining of amyloid is obtained with relatively fresh tissues; long preservation in alcohol or formol tends to weaken the reactions. Probably the best effects are obtained by formol-fixation for 24 hours, and sectioning on the freezing-microtome. Good results may be produced, however, after any of the ordinary fixing and hardening methods by cutting the sections on the freezing-microtome, without imbedding, or imbedding in paraffin. The metachromatic reactions are not satisfactory with celloidin sections. With hæmatoxylin and eosin the amyloid substance stains a light red or bluish-pink; Van Gieson’s stains it a yellow or brownish-pink color, giving it practically the same color that it does epithelial hyalin. The different tissue-relations of the two substances serve to distinguish them. The most important specific amyloid stains are:—

1. Iodine.

1. Stain in Lugol’s solution 5-10 minutes.

2. Dehydrate in absolute alcohol 4 parts, tincture of iodine 1 part.

3. Clear and mount in origanum oil. Seal preparation with paraffin, gold size or shellac.

The iodine reaction is also applied to fresh tissues by pouring Lugol’s solution over a freshly cut surface, and is a good gross test for amyloid. In both microscopic and macroscopic preparations iodine gives a mahogany brown color to amyloid; other tissue is yellow. The iodine reaction may be intensified by placing the sections in a 1 per cent sulphuric acid; the brown color may be changed to blue, violet or green.

2. Methyl Violet.

1. Stain in 0.5 per cent methyl-violet solution ½ to several minutes. Examine in water.

2. Wash in water.

3. Differentiate in 2 per cent acetic or dilute hydrochloric acid 1-2 minutes.

4. Wash thoroughly in water.

5. Mount in lævulose or glycerin-gelatin. Amyloid ruby red; tissue blue-violet.

3. Gentian Violet.

Use same method as for methyl violet. The same color-effects are produced.

4. Methyl Green.

Use methyl green in the same way as methyl violet. Amyloid sky-blue or violet; tissue is green.

5. Iodine Green.

1. Stain for 24 hours in a ⅓ per cent water solution of iodine green.

2. Wash in water.

3. Mount in lævulose, glycerin or glycerin-gelatin. Amyloid red violet; tissue green.

6. Birch-Hirschfeld’s Method.

1. Stain in a 2 per cent alcoholic solution of Bismarck brown for 5 minutes.

2. Wash in absolute alcohol.

3. Wash in water.

4. Stain in a 2 per cent water solution of methyl-violet (or a 20 per cent gentian violet) for 5 minutes.

5. Differentiate in 1 per cent acetic acid until the non-amyloid parts are brown.

6. Wash thoroughly in water.

7. Mount in lævulose, glycerin or glycerin-gelatin. Nuclei are brown; amyloid ruby red.

7. Green’s Method.

To a few cc. of hæmalum in a watch-glass add a saturated solution of methyl-violet, drop by drop, until the mixture shows a faint purple-red tinge at the edge of the glass.

1. Stain sections 15-30 minutes.

2. Differentiate in acid alcohol until the purple begins to fade.

3. Wash thoroughly in water.

4. Mount in glycerin. (Sections may be blotted and dehydrated in pure liquid paraffin; the latter is then removed by blotting with xylol, and then mount in pure white vaseline.)

Nuclei are blue, amyloid ruby-red.

8. Kresyl-echt-violett (Morse’s Method).

Kresyl-echt-violett (R. extra) 1 grm., 5 per cent carbolic acid 80 cc., alcohol 20 cc. Mix, stirring well; filter. Solution keeps well, and can be diluted as desired without precipitating.

1. Stain 1-5 minutes.
2. Wash thoroughly in distilled water, differentiating, if necessary.
3. Blot with filter paper.
4. Dehydrate in absolute alcohol as quickly as possible.
5. Clear in turpentine. Blot nearly dry before mounting.
6. Mount in balsam.

Formol, mercuric chloride and Zenker’s all give good results. Paraffin imbedding, with staining of sections on the cover-glass (albumin-fixative method), is the best method of staining for permanent mounts, although good preparations can be obtained by the use of the freezing-microtome. Carbol-xylol cannot be used for clearing. Nuclei are blue, protoplasm pale blue, amyloid ruby-red.

As a specific reaction for amyloid and mucin this method has been used in my laboratory for the last ten years in preference to any other. The stains are permanent if not exposed to the action of light.

Thionin, toluidin-blue, polychrome-methylene-blue, and other metachromatic dyes are also used to give similar reactions with amyloid, but are not as satisfactory as the kresyl-echt-violett method. Amyloid may also be stained with scarlet R or sudan III, according to the method of Herxheimer, but the results are rarely satisfactory.

II. ATROPHY. Good pictures of atrophic tissues are obtained with formol-Müller’s, mercuric chloride or Zenker’s fixation, and staining with Van Gieson’s, to bring out the stroma which is usually relatively or absolutely increased. In the case of pigment-atrophy the sections should be very thin and stained with alum- or lithium-carmine.

III. CALCIFICATION. Deposits of lime-salts appear in fresh tissue as gritty, refractive areas that are bright and shining by reflected light, and dark by transmitted. They are soluble in acids, solution of the carbonate being accompanied by the formation of bubbles of carbonic acid gas. Hæmalum and the alum-hæmatoxylins show a specific reaction with the phosphates and carbonates of lime, giving them a deep blue or reddish-violet stain. Fresh calcification usually stains diffusely blue; older deposits are deep-blue about the borders of the deposits, lighter or unstained in the center of the mass. Tissues containing much calcium must be decalcified before imbedding. If the process of decalcification is not carried too far the specific staining reaction is not lost.

v. Kossa’s Silver Method for Calcium Phosphate.

1. Fix in alcohol or formol; imbed; cut.

2. Place section in 1-5 per cent silver nitrate solution, and expose to daylight 5 minutes to 1 hour.

3. Wash in distilled water.

4. Transfer section to a 5 per cent solution of sodium hyposulphite, to remove excess of silver nitrate.

5. Wash thoroughly in water.

6. Dehydrate in absolute alcohol.

7. Clear in xylol; mount in balsam.

Calcareous deposit black, as the result of the formation of phosphate of silver and its reduction by the action of light. Alum carmine may be used as a nuclear stain before the sections are treated with silver nitrate, or safranin may be used after the sodium sulphite has been washed out.

IV. CELL GRANULES AND CELL INCLUSIONS. The granules and cell-inclusions here included fall within the class of special protoplasmic structures found particularly in neoplasms and inflamed tissues, and which have been supposed to be parasites. For the staining of other cell-granules see Blood and Blood-forming organs.

1. Altmann’s Granules.

1. Fix small, thin pieces of fresh tissue in equal parts of 5 per cent potassium bichromate and 2 per cent perosmic acid for 24 hours. Wash in running water for several hours. After-harden in alcohol, and imbed in paraffin. Cut very thin and mount on cover-glass; remove paraffin.

2. Stain in aniline-water-acid-fuchsin (acid fuchsin 20 grms., aniline water 100 cc.), warming until vapor is given off.

3. When cool remove the fuchsin with a mixture of 1 part saturated alcoholic picric acid and 2 parts of water.

4. Renew the picric acid solution and warm on the paraffin oven for 30-60 seconds.

5. Dehydrate in alcohol; clear in xylol; mount in balsam.

Protoplasm yellow: Altmann’s granules red: fat black.

2. Russell’s Bodies.

1. Fix and harden in Müller’s; wash; after-harden in alcohol; imbed in paraffin; mount on cover-glass.

2. Stain sections in a saturated solution of fuchsin in 2 per cent carbolic acid 10 minutes or longer.

3. Wash in water.

4. Wash in absolute alcohol for 30 seconds.

5. Counterstain in iodine green (1 grm. in 100 cc. of a 2 per cent carbolic acid) for 5 minutes.

6. Dehydrate quickly in absolute alcohol.

7. Clear in xylol; mount in balsam.

Nuclei are green; Russell’s fuchsin-bodies light-red; Altmann’s granules light-red.

3. Pianese’s Method.

1. Fix in Pianese’s solution (see methods of fixation) 6 hours; wash in running water for 12 hours; after-harden in graded alcohols; imbed in paraffin; mount on cover-glass.

2. Stain 30 minutes in a staining mixture consisting of malachite green 0.5 grm., acid fuchsin 0.1 grm., Martius yellow 0.01 grm., distilled water 150 cc., 96 per cent alcohol 50 cc.

3. Dehydrate in absolute alcohol; clear in xylol; balsam.

Nuclei are green; protoplasm reddish; cell-inclusions light-red.

4. Method for Staining “Plimmer’s Bodies.”

1. Fix in Hermann’s fluid for 12-24 hours. Imbed in paraffin. Mount sections on cover or slide.

2. Transfer sections to hydrogen peroxide for 15-30 seconds.

3. Wash in water.

4. Transfer to a 4 per cent ferric alum solution for 2 hours.

5. Wash in water.

6. Stain in 0.5 per cent watery hæmatoxylin solution for 30 minutes. Differentiate in the ferric alum solution until the nuclei are dark and protoplasm colorless; control under the microscope.

7. Wash in water 3-6 hours.

8. Counterstain in 1 per cent solution of Ehrlich’s neutral red until section is yellow-red.

Nuclei blue-black; cell-inclusions yellow- to copper-red.

V. CHOLESTERIN. Cholesterin is soluble in absolute alcohol, xylol, ether and glacial acetic. It occurs in the tissues in characteristic rhombic plates often showing a square notch in one corner. In sections from which the cholesterin has been dissolved its presence may be told by the appearance of “cholesterin-clefts” in the tissue, or often in the protoplasm of large foreign-body giant-cells (“cholesterin-giant-cells”). With concentrated sulphuric acid, sections or material containing cholesterin become yellow and then rose-pink. Lugol’s gives it a brown color which turns blue-violet after the addition of sulphuric acid, and exhibits a play of colors, blue, green, to red.

VI. CLOUDY SWELLING. This is best seen in the fresh state in cells obtained by scraping or teasing, or by the examination of frozen sections. Osmic acid, sudan III, scarlet R. ether, alcohol and acetic acid may be used to differentiate from fatty degeneration. The ordinary fixing and staining methods give good pictures, except for slight degrees of the change. These are sometimes wholly lost as the result of the contraction due to the fixation.

VII. COLLOID. (See Epithelial Hyalin.)

VIII. CORNIFICATION. Horn takes the plasma stains (eosin, picric acid, etc.). Van Gieson’s makes a good differential stain. With Gram’s method horn stains deep blue, and with the Ehrlich-Biondi-Heidenhain method it stains red. After fixation in Flemming’s it may be stained with safranin or gentian-violet. Keratohyalin occurs as fine granules in the cells of the stratum granulosum. They stain by hæmatoxylin, carmine and Gram’s method, or may be demonstrated by means of special stains. Eleidin stains with carmine and the fat-stains, but not with hæmatoxylin.

1. Buzzi’s Method of Staining Eleidin and Keratohyalin.

1. Harden, imbed, cut.

2. Stain in Congo red (2-3 drops of a 1 per cent water solution added to small basin of water) for 2-3 minutes.

3. Wash thoroughly in water.

4. Stain in hæmatoxylin, and wash.

5. Dehydrate in absolute alcohol; xylol; balsam.

Keratohyalin blue, eleidin red.

2. Fick’s Method of Staining Keratohyalin and Keratin.

1. Harden in alcohol, imbed, cut.

2. Stain in saturated water solution of kresyl-echt-violett for 3-4 minutes.

3. Wash thoroughly in water.

4. Differentiate in 95 per cent alcohol until connective-tissue is colorless.

5. Dehydrate in absolute alcohol; xylol; balsam.

Keratohyalin red, keratin dark violet; nuclei blue-violet, plasma light blue-violet.

IX. FAT. When alcohol has been used in the preparation of the tissue, the fat-contents of the latter are dissolved out, and their presence can alone be told by the presence of vacuoles. When osmic acid is used as a fixing agent the oleates and oleic acid are blackened. The tissue should then be washed in running water and cut upon the freezing-microtome, or it may be imbedded in celloidin or paraffin if this is done as quickly as possible to prevent the loss of the fat. Chloroform or benzene should be used in place of xylol, as the last-named dissolves out the fat. Safranin should be used as a stain after fixation with any fluid containing osmic acid. Frozen sections are to be mounted in glycerin-gelatin; when balsam is used it should be warm melted Canada balsam without xylol. Formol fixation preserves fat, and tissues so fixed may be cut on the freezing-microtome and the sections stained with osmic acid, sudan III or scharlach R, with nuclear counterstaining when desired. For the demonstration of fat-embolism, fatty degeneration or fatty infiltration the following methods are advised:—

1. Staining of Fat with Osmic Acid.

1. Fix in formol for 24 hours.

2. Wash; freeze; cut.

3. Place sections in 1 per cent osmic acid, Flemming’s or Marchi’s fluid for 1-24 hours.

4. Wash in water, changing frequently.

5. 80 per cent alcohol ½-2 hours.

6. Wash in water.

7. Place section flat on slide; blot; add a drop of warmed glycerin-gelatin; cover quickly. Ringing or sealing is not necessary.

Or, to mount section in balsam:—

After 6, counterstain with hæmatoxylin or safranin; wash again; dehydrate quickly with absolute alcohol; clear in pure benzene; mount in pure melted Canada balsam (containing no xylol).

2. Staining of Fat with Sudan III or Scharlach R.

Staining-solutions of these dyes may be made, as follows:—

a. Dissolve stain in 70-80 per cent boiling alcohol, keep in the incubator over night, and use warm.

b. Make a solution of absolute alcohol 70 cc., 10 per cent caustic soda solution 20 cc., water 100 cc. Saturate this with the stain, slightly heating.

c. Make a mixture of 70 per cent alcohol 50 cc. and pure acetone 50 cc.; saturate this with the stain.

All solutions of these dyes should be filtered before using, and should be kept covered to avoid evaporation and subsequent precipitation.

1. Formalin fixation 24 hours; cut on freezing-microtome.

2. Place sections in 70 per cent alcohol.

3. Stain in the simple solution 20-30 minutes; in the acetone or alkaline alcoholic solutions 2-3 minutes.

4. Wash in 50-70 per cent alcohol, differentiating as needed.

5. Transfer to water; thence to slide; blot, and mount in glycerin gelatin.

When a nuclear counterstain is desired, put the sections in water after 4; then stain in hæmatoxylin; differentiate quickly in acid alcohol; wash in water; place in weak ammonia or lithium-carbonate solution; wash in water; transfer to slide; blot; mount in glycerin gelatin.

Sudan III and scarlet R stain the smallest particles of fat yellowish-red to deep scarlet; scarlet R on the whole gives the best results. The contrast with the blue nuclei when stained with hæmatoxylin gives beautiful preparations.

3. Staining of Fat with Indophenol.

Stain sections with lithium-carmine; wash; then stain 20 minutes in a saturated solution of indophenol in 70 per cent alcohol. Fat blue; nuclei red.

4. Staining of Fatty Acids and Soaps.

a. Benda’s Method. Fix in 10 per cent formol. Transfer tissue to Weigert’s copper-fluorchrom mordant (neutral copper acetate 5 grms., fluorchrom 2.5 grms., water 100 cc.; boil and add 5 cc. of 36 per cent acetic acid) in the incubator for 2-4 days. Cut on the freezing-microtome. Stain sections in sudan III or scharlach R, and then in hæmatoxylin. Nuclei are blue, normal fat red, necrosed fat green due to formation of fatty acid copper salt. Soaps give the same reaction when converted into insoluble salts by fixing in formol saturated with calcium salicylate. Through comparison of tissue hardened in this way with another portion fixed in formol alone soaps and fatty acids may be differentiated.

b. Smith’s Method. Stain in concentrated water solution of Nile blue sulphate for 10 minutes. Fat stains red, nuclei dark blue, protoplasm light blue, fatty acids dark-blue. Differentiate in 1 per cent acetic acid; wash in water; mount in glycerin-gelatin.

X. FIBRIN. Fibrin stains with the acid aniline dyes, except in areas of necrosis containing diffused chromatin, under which conditions it stains deep blue with hæmatoxylin. In Van Gieson’s mixture it stains yellow or brownish; in Mallory’s reticulum stain it stains red, and with Mallory’s chloride of iron hæmatoxylin it is grayish to dark blue. The best selective method by far is Weigert’s, and it is the only really practical method giving a good differentiation.

1. Weigert’s Fibrin Stain.

I have obtained the best results by making this stain as follows: 10 cc. of aniline oil and 100 cc. of water are shaken together violently for several minutes, and then filtered through a moist filter. The filtrate must contain no drops of aniline. Add to the filtrate sufficient dry gentian-violet or methyl-violet to produce a metallic shimmer on the surface of the solution after the dye is dissolved by shaking. The solution will keep for several months.

Weigert advised the use of two stock solutions, I (absolute alcohol 33 cc., aniline oil 9 cc., methyl violet in excess) and II (saturated water solution of methyl violet). These solutions will keep for years. When ready to use stain take 3 cc. of Sol. I and 27 cc. of Sol. II. This staining mixture will keep for about 2 weeks.

1. Fix in alcohol, formol, acetone, mercuric chloride or Müller’s. Imbed in celloidin or paraffin; the latter preferably. Mount sections on cover-glass with albumin fixative. Celloidin sections must be fastened to slide by thin film of celloidin to prevent shrinkage. Sections fixed in chromic mixtures (and sometimes after formol fixation) must be oxidized in potassium permanganate and then reduced in oxalic acid to give good results. (Transfer sections to a 1 per cent solution of potassium permanganate to which 2 volumes of water have been added; oxidize for 10 minutes; then wash in water, and reduce for several hours in a 5 per cent water oxalic acid solution.)

2. Wash in water.

3. Stain in lithium carmine; differentiate in acid alcohol; wash thoroughly in water.

4. Stain on the slide or cover-glass in the aniline-methyl-violet (or gentian-violet) solution for 10 minutes. Wash off stain with physiologic salt solution.

5. Blot section with absorbent paper.

6. Cover section with Lugol’s (300-2-1) or a 5 per cent watery potassium iodide saturated with iodine. Leave on section for 1-5 minutes.

7. Blot off iodine.

8. Differentiate in aniline xylol (equal parts of xylol and aniline oil) until the nuclei become red.

9. Wash in xylol, blotting with absorbent paper. Repeat until section is transparent; then mount in balsam. All aniline oil must be removed before using the balsam.

Nuclei are red; fibrin deep blue; bacteria, mucin, keratin and Altmann’s granules also blue. The differentiation must be carefully controlled under the microscope, and should be stopped before the finest threads of fibrin begin to be decolorized.

XI. GLYCOGEN. Glycogen is soluble in water; and fixation and hardening must be carried out with absolute alcohol to prevent the solution of the glycogen. Tissue must be fixed immediately after death, as glycogen is quickly broken up. Its reaction with iodine is similar to that of amyloid, but it does not give the iodine-sulphuric-acid reaction that the latter substance does.

1. Best’s Iodine Method.

1. Fix and harden in absolute alcohol; imbed in paraffin; cut.

2. Stain somewhat deeply with hæmatoxylin.

3. Wash in water.

4. Stain in iodine 1, potassium iodide 2, water 100.

5. Dehydrate in iodine 2, absolute alcohol 100.

6. Differentiate in origanum oil, 1-2 hours.

7. Wash thoroughly with xylol.

8. Arrange on slide and dry in air.

9. Mount in pure melted balsam (no xylol).

Nuclei are blue, glycogen brown.

2. Best’s Carmine Method for Glycogen.

1. Fix in absolute alcohol; imbed in celloidin; cut.

2. Stain in hæmatoxylin; differentiate in acid alcohol.

3. Wash in water.

4. Stain in filtered carmine mixture (carmine 1 grm., ammonium chlorate 2 grms., lithium carbonate O.5 grm., water 50 cc.; bring to boiling point, and when cool, add 20 cc. of strong liquid ammonia. Keep in dark; can be used after 2-3 days and gives good results up to 14 days) 2 parts, strong ammonia 3 parts, methyl alcohol 6 parts. Make fresh each time it is used, as it soon precipitates; do not filter; stain few sections at a time ¾-1 hour.

5. Differentiate 1-2 minutes in a mixture of absolute alcohol 4 parts, methyl alcohol 2 parts, water 5 parts.

6. Wash in 80 per cent alcohol.

7. Dehydrate in absolute alcohol.

8. Clear in xylol; mount in balsam.

Glycogen is stained red; nuclei blue; dense connective-tissue, mast-cell granules, protoplasm of gastric glands, etc., red; but these can all be distinguished morphologically from glycogen. This is by far the best method for the staining of glycogen.

XII. HYALIN. Epithelial hyalin (colloid) stains red or violet with hæmatoxylin and eosin; it takes the other acid dyes and stains to some degree with basic aniline stains. Van Gieson’s stains it a yellow, orange or brownish-pink. Kresyl-echt-violett gives it a deep indigo-blue color or a more green robin-egg blue. Connective-tissue hyalin stains deep brilliant red with Van Gieson’s; this is the best method for differentiating connective-tissue hyalin from amyloid or epithelial hyalin. Russell’s method also stains hyalin red.

XIII. HYDROPIC DEGENERATION. Fix by heat or formol-alcohol. Imbed in celloidin; stain with hæmatoxylin and eosin.

XIV. HYPERTROPHY. Fix in Müller’s or mercuric chloride for simple staining; for study of nuclei fix in Flemming’s and stain with safranin.

XV. INFLAMMATION. The process of inflammation may be studied to advantage in the mesentery, web or tongue of the curarized living frog, by stretching these parts over a cork-ring attached to a glass plate on which the animal rests. The exposed tissues must be kept moist with physiologic salt solution. Heat, chemicals or other irritants may be employed to produce the inflammatory reaction. For the study of the inflammatory process in sections the ordinary fixations may be employed, but for the study of the nuclei, mitotic figures and cell-granulations fixation in Flemming’s, Zenker’s, etc., is advised. Safranin, methylene blue and eosin, the various stains used in the study of blood-cells, etc., may be used.

1. Staining of Mast-cells.

a. Kresyl-echt-violett used as for amyloid or mucin is the best stain for mast-cells. The cell-granules stain bright rose-red.

b. Ehrlich’s Dahlia Method.

1. Harden in absolute alcohol; imbed; cut.

2. Stain with saturated water solution of dahlia.

3. Wash in water.

4. Dehydrate in absolute alcohol.

5. Clear in xylol; mount in balsam.

c. Unna’s Method for Mast and Plasma Cells.

1. Harden in absolute alcohol; imbed; cut.

2. Stain in Unna’s polychrome methylene blue ¼-12 hours.

3. Wash in water.

4. Differentiate in Unna’s glycerin-ether mixture (Grübler) 15 seconds to several minutes.

5. Wash carefully in water.

6. Dehydrate in absolute alcohol; clear in xylol; mount in balsam.

Mast cell granules are red; plasma cell granules blue.

d. Various modifications of the Romanowsky method stain mast- and plasma-cells very well.

XVI. IODINE. For the demonstration of iodine in tissues the following method has been advised by Justus. The experience of other workers with it has not been satisfactory.

1. Harden in absolute alcohol; imbed in celloidin; cut.

2. Soak in water to remove alcohol.

3. Put section in wide-mouthed, stoppered bottle in freshly prepared green chlorine-water for 1-2 minutes.

4. Transfer section on a glass needle to a vessel containing 500 cc. water and 1 cc. of a 1 per cent solution of silver nitrate for 2-3 hours. The section is colored yellow-green, and a precipitate of silver chloride appears.

5. Transfer section to a warm saturated solution of sodium chloride until it becomes light.

6. Wash in distilled water.

7. Transfer to a concentrated solution of mercuric chloride.

8. Examine in pure glycerin.

Iodine should be red.

XVII. MITOTIC FIGURES. Various histologic methods devised for the study of mitoses can be applied to the demonstration of these in neoplasms, inflammation and regeneration. Flemming’s solution or mercuric chloride fixation gives best results, although formol, or even absolute alcohol, when used quickly and carefully gives fair results if tissue is very fresh.

1. Flemming’s Solution and Safranin.

1. Fix small pieces of fresh tissue in Flemming’s, in the dark, for 24 hours; wash 24 hours; after-harden in graded alcohols; imbed and cut.

2. Stain in 1 per cent water solution or saturated aniline water solution of safranin, or 1 per cent water methyl violet for 12-24 hours, or carbol-fuchsin for one hour.

3. Differentiate quickly in a 0.5-0.0001 HCl in 70 per cent alcohol and then in absolute alcohol until stain no longer comes away in clouds and nuclei have right shade.

4. Clear in xylol; mount in balsam.

Fat is black; mitoses stand out sharply; tubercle-bacilli may be stained black or red.

2. Fixation in mercuric-chloride may be followed by Ehrlich-Biondi-Heidenhain’s stain (saturated aqueous orange 100 cc., saturated aqueous acid fuchsin 20 cc., saturated aqueous methyl green 50 cc.) 12 grms. of Grübler’s prepared stain dissolved in 100 cc. of distilled water, for stock solution. For staining take 1 cc. of stock solution, water 30 cc., ½ per cent watery acid fuchsin 3 cc., and 2 per cent acetic 5-6 drops. Stain 2-24 hours; wash in 90 per cent alcohol; dehydrate in absolute; clear in xylol; balsam.

Resting nuclei are bluish; mitoses and fragments of leukocyte nuclei dark green; red blood cells orange red; protoplasm and connective-tissue fuchsin red.

3. Benda’s Iron-Haematoxylin Method.

1. Fix in osmic acid, mercuric chloride or other fixative.

2. Stain sections by placing them in liq. ferri. sulfur. oxyd. (Germ. Pharm.) diluted with double its volume of water, for 24 hours; wash carefully in distilled water and then in tap water; stain in 1 per cent watery hæmatoxylin until section is black. Wash in water. Differentiate in 10-30 per cent acetic acid, or in liq. ferri. sulfur. oxyd. diluted with distilled water 1-20. A 10 per cent solution of ferric sulphate may be used instead of the persulphate.

4. Heidenhain’s Iron-Haematoxylin.

1. Imbed in paraffin after fixation in mercuric chloride.

2. Immerse section in a 1.5 per cent solution of iron-alum sulphate (violet-colored salt) or iron-ammonium sulphate for ½-3 hours.

3. Wash in water.

4. Stain in 0.5 per cent watery hæmatoxylin or hæmatein for 12-18 hours.

5. Wash in water.

6. Differentiate in the iron-alum or iron-ammonium solution until the section becomes deep blue (control under microscope) and nuclear structures stand out distinctly.

7. Wash in running water for 15 minutes.

8. Absolute alcohol; xylol; balsam.

Instead of the watery hæmatoxylin solution a mixture of hæmatoxylin 1 grm., alcohol 10 cc., and water 90 cc. may be used. Keep four weeks before using. Stain 24-36 hours. For contrast staining a weak solution of Bordeaux red may be used before the iron-alum and hæmatoxylin, staining 24 hours.

XVIII. MUCIN. Mucin stains a deep blue or reddish-violet with an over-ripe hæmatoxylin. When counterstained with picric acid very beautiful preparations can be obtained. Mucin also gives a metachromatic reaction with kresyl-echt-violett, thionin, toluidin-blue and polychrome methylene-blue, staining red with these stains. Water or carbolic-acid solutions of these stains may be used; dehydrate in absolute alcohol, clear in xylol, and mount in balsam. In my opinion Morse’s Carbol-kresyl-echt-violett method as given above for amyloid is the best of these metachromatic reactions. Muchæmatein and mucicarmin give the most delicate reactions.

1. Mayer’s Muchaematein.

1. Absolute alcohol fixation is preferable.

2. Stain sections in Mayer’s solution (hæmatein O.2 grm. mixed with a few drops of glycerin, O.1 grm. of aluminum chloride, 40 cc. of glycerin, 60 cc. of water) for 5-10 minutes.

3. Wash in water.

4. Dehydrate in absolute alcohol; xylol; balsam.

Carmine may be used for counterstaining; mucin is blue. Should the mucin swell in the stain replace water and glycerin with 100 cc. of 70 per cent alcohol and 1-2 drops of nitric acid.

2. Mayer’s Mucicarmin.

Make staining solution by mixing 1 grm. carmine, O.5 grm. aluminum chloride, 2 cc. water and 100 cc. of 50 per cent alcohol, heating over the flame for 2-3 minutes until mixture darkens. Let stand 24 hours and filter. The stock solution may be diluted 1-10. Stain 10 minutes. If it does not stain well add 0.5-1 grm. of aluminum chloride. Mucin alone should be stained red. Counterstain with hæmatoxylin.

XIX. MYELIN. This appears in the form of doubly refractive granules, that stain with less intensity with the fat dyes, but may be differentiated from fat in that it loses the power of reducing osmic acid after being mordanted for eight days or more in bichromate solutions, while fat does not.

XX. NECROSIS. Hæmatoxylin and eosin, and Van Gieson’s give good pictures. Use Weigert’s fibrin stain for coagulation-necrosis, and Benda’s method for the demonstration of fatty acids for the staining of fat-necrosis. Recent necrotic areas stain diffusely blue with hæmatoxylin; older areas may take the plasma stains alone. Use various methods for the demonstration of micro-organisms in the necrotic areas.

XXI. NEOPLASMS. Use hæmatoxylin and eosin, and Van Gieson’s for ordinary diagnosis. To differentiate sarcoma and carcinoma use Van Gieson’s, Mallory’s or other reticulum stains. For the study of cell-inclusions use Altmann’s, Russell’s, Plimmer’s and Pianese’s methods. Special fixation or Zenker’s is necessary. Methylene-blue and eosin after Zenker’s give excellent pictures. For the demonstration of mitoses the methods given above should be employed.

XXII. PIGMENT. Use the carmines for contrasting melanin, hæmofuscin, lipochromes, hæmatoidin, hæmosiderin, bilirubin and all yellow, brown, blue, black, etc., extrinsic pigments. In tissue fixed in mercuric chloride or formol bilirubin is green, and can thus be differentiated from hæmatoidin. The lipochromes give weak fat-reactions, and this is used to distinguish them from other yellow or brown pigments. Alcohol fixation is the best for pigment study, although the other fixing solutions may be used. Formol sometimes produces pseudo-pigments by its action upon hæmoglobin. The iron-reactions are obtained best in sections cut on the freezing-microtome, although both paraffin and celloidin imbedding may be used. In testing for iron glass needles should be used and all traces of iron should be removed from staining-dishes, slides, etc., by treating with hydrochloric acid, distilled water and alcohol.

1. Potassium Ferrocyanid Test for Iron.

1. Stain sections in lithium carmine for several hours.

2. Differentiate in acid alcohol, stopping short of the desired complete differentiation of the nuclei.

3. Wash in water.

4. Saturated solution of potassium ferrocyanid 1-3 hours.

5. Acid alcohol until iron-pigment becomes blue (½-12 hours). Complete differentiation of nuclei.

6. Wash in water.

7. Dehydrate in absolute alcohol.

8. Clear in xylol; mount in balsam.

Hæmosiderin is blue (Berlin blue); nuclei are red. Lithium carmine may be used after the iron-test, if desired.

2. Ammonium Sulphide Test for Iron.

1. Fix in alcohol; imbed; cut.

2. Treat sections with yellow ammonium sulphide for 5-60 minutes.

3. Wash quickly in water.

4. Dehydrate in absolute alcohol.

5. Clear in xylol; mount in balsam.

Stain with lithium carmine either before or after the reaction with ammonium sulphide. Iron is grayish-black to black.

3. Combined or Masked Iron.

1. Treat tissues with Bunge’s fluid (95 per cent alcohol 95 cc., 25 per cent hydrochloric acid 10 cc.) for 1-2 hours at 50-60°C., until inorganic iron is all removed.

2. Place tissues in acid alcohol (sulphuric acid 4 cc. in 100 cc. alcohol 95 per cent.)

5. Wash sections in acid alcohol, then pure alcohol, and finally in distilled water.

6. Transfer to ammonium sulphide (5-60 minutes) or to potassium ferrocyanid and O.5 HCl for 5 minutes.

7. Wash in water.

8. Counterstain in eosin or safranin; wash; dehydrate in absolute alcohol; clear in cedar-oil; mount in benzene balsam. Keep preparations in the dark.

4. Staining of Chromophilic Cells.

1. Fix in a chromic solution. In this the chromophilic cells become yellow or brown.

2. Stain in polychrome methylene blue; the cells become grass-green in color.

5. Tests for Silver, Lead and Mercury.

Use ammonium sulphide as for iron. Black sulphides are formed.

6. Test for Copper.

Treat with potassium ferrocyanid and hydrochloric acid; copper gives a dark yellow-brown coloration.

XXIII. PSEUDOMUCIN. It is not precipitated by acetic acid. It has a greater affinity for the diffuse stains than mucin, and gives weaker metachromatic reactions.

XXIV. REGENERATION AND REPAIR. For the staining of mitoses, cell granules and cell-inclusions see methods given above. See also methods for staining of epithelium, reticulum, neuroglia, etc.

XXV. URIC ACID AND PURIN BASES:—

1. Courmont and Andre’s Method.

1. Fix in absolute alcohol; imbed; cut.

2. Treat sections with 1/100 ammonia solution or very weak sodium hyposulphite solution.

3. Transfer to 1/100 silver nitrate solution.

4. Wash.

5. Develop with a photographic developer.

6. Wash in water; stain with hæmalum and eosin; dehydrate; clear in xylol; balsam.

Uric acid and xanthin or purin bases appear as black granules.

CHAPTER XXVII.
THE STAINING OF PATHOGENIC MICRO-ORGANISMS IN TISSUES

Rapid fixation and hardening are requisites for the successful staining of micro-organisms in sections. Alcohol, Zenker’s, mercuric chloride and formol give best results; Müller’s because of its slow action is not good, although formol-Müller’s may be used because of the more rapid fixation with this fluid. In the case of formalin-fixation staining with Weigert-Gram’s method may not give good results unless the sections are oxidized in potassium permanganate solution and then reduced in oxalic acid. (See Staining of Fibrin.) Preservation of the tissue for a long time in alcohol impairs the staining power of micro-organisms contained within it. The tissue should be imbedded preferably in paraffin, as very thin sections must be obtained. The freezing-microtome may be employed and the thinnest sections selected for staining. Celloidin stains very heavily with the aniline dyes and retains the color, so that bacteria in celloidin sections do not stand out very distinctly. On the whole paraffin sections, floated on slide or cover, and fastened by albumin-fixative, give the best results, though for the micro-organisms stained in carbol-fuchsin and decolorized in nitric acid it is best to float the sections directly onto the warm stain without removing the paraffin, and mount without the use of alcohol. This method may be employed for all stains that are taken out by alcohol. The stains used for film preparations are as a rule applicable to sections. The basic aniline dyes, particularly methylene-blue, fuchsin, methyl or gentian violet, kresyl-echt-violett, thionin, and Bismarck brown, either in saturated alcoholic solutions or dilutions of such, or in combination with alkalies, aniline oil or phenol, are usually employed. The various modifications of the Romanowsky method are very useful. The time required for staining in sections is usually much longer than for films; but the staining can often be accelerated or strengthened by warming over the flame or in the incubator. Contrast staining of the nuclei with lithium-carmine or Bismarck brown is advisable after the use of staining methods in which the nuclei are decolorized. Xylol or origanum oil should be used for clearing.

I. THE STAINING OF BACTERIA IN TISSUES.

According to their staining-reactions bacteria may be very conveniently grouped in three classes: 1, Staining with Gram-Weigert’s method; 2, Not staining with Gram-Weigert’s; 3, Staining with the tubercle-bacillus method (acid-resisting).

1. BACTERIA STAINING BY THE GRAM-WEIGERT METHOD.

Weigert’s modification of Gram’s method, as given above for the staining of fibrin, is the best for the staining of bacteria that stain by this method. (See Fibrin, Chapter [XXVI].) The differentiation with aniline-xylol is slower and safer than with alcohol. Acetone-xylol (1:5) has been recommended in place of aniline-xylol. Wolbach recommends the use of a 5-10 per cent colophonium-alcohol for differentiation. Contrast staining with watery Bismarck brown, dilute carbol-fuchsin or eosin may be carried out if desired. The aniline-xylol may be saturated with eosin and the section stained during the differentiation. Carbol-gentian-violet may be used instead of aniline-gentian-violet; it keeps much better than the latter.

Staining by Gram’s Method (Gram-positive).

Staphylococcus pyogenes aureus.
Staphylococcus pyogenes albus.
Staphylococcus pyogenes citreus.
Streptococcus pyogenes.
Micrococcus tetragenus.
Diplococcus pneumoniæ.
Bacillus aërogenes capsulatus.
Bacillus of diphtheria.
Bacillus of anthrax.
Bacillus of leprosy.
Bacillus of tetanus.
Bacillus of tuberculosis.
Bacillus of rhinoscleroma.
Bacillus of mouse septicæmia.
Bacillus of swine erysipelas.
Oïdium albicans.
Mycelium of actinomyces.

2. BACTERIA NOT STAINING BY GRAM’S METHOD.

For the bacteria belonging to this class Löffler’s methylene-blue, carbol methylene-blue, a watery solution of methylene-blue or gentian-violet, Leishman’s or Wright’s modification of Romanowsky’s methylene-blue eosin method (see page [290]), Unna’s alkaline methylene-blue solution preceded by eosin after Zenker’s fixation (see page [260]), aniline gentian-violet, Zieler’s method and carbol fuchsin are most commonly used as stains. Wolbach advises the use of a 5-10 per cent acetone-colophonium solution for the differentiation of Gram-negative bacteria in tissue fixed in formol.

1. Löffler’s Methylene-blue.

1. Saturated alcoholic solution of methylene-blue 30 cc.; potassium hydrate solution (1 in 10,000) 100 cc.

2. Stain 5 minutes to 24 hours.

3. Wash in water.

4. Differentiate in 1 per cent acetic acid, 10-30 seconds.

5. Wash in 90 per cent alcohol, 2-5 minutes; dehydrate in absolute alcohol; clear in xylol; mount in balsam.

2. Gentian-violet.

1. Stain sections in a 2 per cent watery gentian-violet for 5-20 minutes.

2. Wash in water.

3. Decolorize in 70 per cent alcohol until stain ceases to come away.

4. Dehydrate in absolute alcohol; clear in xylol; balsam.

3. Zieler’s Method.

1. Fix in Orth’s solution, or any fixing solution except those containing osmic acid; imbed in paraffin or celloidin.

2. Stain in Pranter’s solution (orcein D 0.1 grm., hydrochloric acid 2.0 cc., 70 per cent alcohol 100 cc.) for 8-24 hours.

3. Wash rapidly in 70 per cent alcohol.

4. Wash in water.

5. Stain in polychrome methylene-blue 10 minutes to several hours.

6. Wash in distilled water.

7. Differentiate in glycerin ether until no more clouds of color come away and section is light blue.

8. Wash in distilled water.

9. 70 per cent alcohol for a few seconds; absolute 5-10 minutes; xylol; balsam.

Protoplasm is gray-brown; bacteria dark-blue; background colorless. Zieler’s method is especially good for the staining of the glanders, typhoid and chancroid bacilli and the gonococcus.

For Unna’s methylene-blue eosin and the modifications of the Romanowsky method see Pages [260] and [290] respectively. Pappenheim’s methyl-green-pyronin method is also recommended for the staining in sections of Gram-negative bacteria.

Not Staining by Gram’s (Gram-negative).

Gonococcus.
Micrococcus melitensis.
Meningococcus (in sections).
Bacillus of bubonic plague.
Bacillus of chancroid.
Bacillus coli communis.
Bacillus dysenteriæ.
Bacillus of epidemic conjunctivitis (Koch-Weeks).
Bacillus of influenza.
Bacillus mallei.
Bacillus pneumoniæ.
Bacillus proteus.
Bacillus of malignant œdema.
Bacillus pyocyaneus.
Bacillus of typhoid fever.
Bacillus of fowl cholera.
Bacillus of rabbit septicæmia.
Bacillus of swine plague.
Spirillum of Asiatic cholera.
Spirochæte pallida.

3. BACTERIA STAINING BY THE TUBERCLE-BACILLUS METHOD. (ZIEHL-NEELSEN.)

1. Tubercle-bacillus.
2. Lepra-bacillus.
3. Smegma-bacillus.
4. Lustgarten’s bacillus.

1. Stain sections by floating thin paraffin sections directly on to warm carbol-fuchsin (fuchsin 1 grm., absolute alcohol 10 cc., cryst. carbolic acid, 5 grms., water 100 cc.) for 1-3 minutes.

2. Transfer on spatula to water, agitating so as to wash off excess of stain.

3. Transfer section to 30 per cent nitric acid and water alternately, until section has a pale lilac tint.

4. Wash in water.

5. Float on warm watery methylene-blue for 1 minute.

6. Wash in water.

7. Float section on slide; dry over flame or in oven; melt over flame, and put section at once into xylol to remove paraffin.

8. Balsam.

Various staining-methods have been recommended for the staining of the most important pathogenic bacteria in tissues. The most useful of these methods are here given:—

a. Cocci.

1. Pyogenic Cocci. Stain by Gram-Weigert’s, contrast with Bismarck brown or lithium-carmine.

2. Pneumococcus. Stains with ordinary water solutions, carbol-fuchsin and Gram-Weigert’s. The staining of the capsule in sections is not very satisfactory.

3. Gonococcus. Gram-negative. Stains in sections with Zieler’s method, Löffler’s methylene-blue, or dilute carbol-fuchsin with differentiation in alcohol.

4. Micrococcus Catarrhalis. Stains like the gonococcus.

5. Diplococcus Intracellularis Meningitidis. Smear preparations often Gram-positive, in sections usually Gram-negative. Use same stains as for gonococcus.

6. Micrococcus Tetragenus. Stain with Gram’s or watery solutions of basic aniline dyes.

b. Bacilli.

1. Anthrax-bacillus. Stain with Gram-Weigert’s and contrast with Bismarck brown or lithium-carmine. Stains also with strong watery gentian-violet solution, with differentiation in strong alcohol.

2. Bacillus of Malignant Oedema. Gram-negative. Stain with watery solution or gentian-violet.

3. Bacillus of Tetanus. Gram-positive. Stains with watery solutions of basic aniline dyes.

4. Bacillus Aërogenes Capsulatus. Gram-positive. Stains with other aniline stains.

5. Bacillus Pyocyaneus. Stains with Gram’s and other aniline dyes.

6. Bacillus of Influenza. Gram-negative. Fix tissue in alcohol. Stain with dilute carbol-fuchsin and differentiate in dilute acetic acid.

7. Koch-Week’s Bacillus. Gram-negative.

8. Bacillus of Bubonic Plague. Gram-negative. Stain by Gaffky’s method (Fix in alcohol or a mixture of glacial acetic acid 10.0, chloroform 30.0, and 96 per cent alcohol 60.0, imbed in paraffin, stain 2-3 hours in weak watery methylene-blue, dehydrate quickly in absolute alcohol, xylol, balsam). It may also be stained by 24 hours in concentrated solution of fuchsin in glycerin, rapid differentiation in weak acetic; alcohol; xylol; balsam. Alcohol or mercuric chloride fixation should be used, as formol fixation does not give good staining.

9. Typhoid Bacillus. Löffler’s methylene-blue or carbol-fuchsin, staining 24 hours, decolorizing in dilute acetic and washing rapidly in alcohol. Zieler’s method may also be used. It is Gram-negative.

10. Paratyphoid Bacillus. Stains like the typhoid bacillus.

11. Colon Bacillus. Gram-negative. May be stained with Löffler’s methylene-blue or carbol-fuchsin.

12. Diphtheria Bacillus. May be stained in sections of diphtheritic membranes with Löffler’s methylene-blue, watery aniline stains, or with Gram’s if the decolorization is not carried too far.

13. Bacillus of Chancroid. Gram-negative. Stain according to Unna’s method:—

1. Fix in alcohol.

2. Stain 5-10 minutes in a mixture of a solution of methylene-blue 1 grm., potassium carbonate 1 grm., alcohol 20 cc., water 100 cc., and a solution of methylene-blue 1 grm., borax 1 grm., water 100 cc.

3. Place sections on slide; blot.

4. Decolorize in Unna’s glycerin-ether mixture.

5. Dry; dehydrate in alcohol.

6. Xylol; balsam.

14. Bacillus of Glanders. Gram-negative. Stain with Zieler’s method or with Löffler’s methylene-blue, differentiating in weak acetic.

Noniewicz’s Method.

1. Stain with Löffler’s methylene-blue 2-5 minutes.

2. Wash in water.

3. Differentiate for about 5 seconds in a mixture of ½ per cent acetic acid 75 cc., ½ per cent watery solution tropaeolin 25 cc.

4. Wash in water; dry by blotting; xylol; balsam.

Bacilli deep-blue; tissues light-blue.

15. Bacillus of Rhinoscleroma. Gram-positive. Fix in alcohol for Wolkowitsch’s method:—

1. Stain in aniline gentian-violet 24-48 hours.

2. Wash in water.

3. Treat with Lugol’s 1-4 minutes.

4. Decolorize in absolute alcohol.

5. Remove more color by oil of cloves.

6. Xylol; balsam.

In tissues fixed in osmic acid and then stained in hæmatoxylin the bacilli are dark blue with light blue capsules. The hyaline substance of rhinoscleroma stains with basic stains.

16. Friedländer’s Bacillus. Gram-negative. Stains with ordinary aniline dyes. For staining the capsules the following method is advised:—

1. Stain for 24 hours in the incubator in a mixture of a concentrated alcoholic solution of gentian-violet 50 cc., glacial acetic acid 10 cc., and distilled water 100 cc.

2. Wash in a 1 per cent acetic acid solution.

3. Alcohol; xylol; balsam.

Bacilli deep-blue; capsules light-blue.

17. Tubercle-bacillus. Gram-positive. Stain in sections on warm carbol-fuchsin without removing paraffin, as given above. Alcohol and mercuric chloride fixation give best results. Aniline-gentian-violet may also be used, staining with a warm solution for 15-30 minutes, and decolorizing in 20 per cent nitric acid followed by 70 per cent alcohol, counterstaining in Bismarck brown, dehydrating in alcohol, clearing in xylol and mounting in balsam. The Weigert-Gram method may be used for the demonstration of the branched or streptothrix forms of the tubercle bacillus. For celloidin sections Mallory and Wright advise the following:—

1. Stain rather lightly in alum-hæmatoxylin.

2. Wash in water.

3. Dehydrate in 95 per cent alcohol.

4. Attach sections to slide by ether-vapor method.

5. Stain in steaming carbol-fuchsin 2-5 minutes.

6. Wash in water.

7. Acid alcohol ½-1 minute.

8. Wash thoroughly in several changes of water to remove acid completely and to bring back blue color to nuclei.

9. 95 per cent alcohol to remove fuchsin.

10. Aniline-oil, followed by xylol, blotting.

11. Xylol; balsam.

Celloidin is colorless, nuclei blue, tissue colorless, tubercle-bacilli red. Orange G may be used as a diffuse stain.

18. Lepra Bacillus. Gram-positive. Stain paraffin sections on warm carbol-fuchsin, as for the tubercle-bacillus. To differentiate from the tubercle-bacillus, stain 6-7 minutes in a dilute alcoholic solution of fuchsin, and decolorize in acid alcohol (nitric acid 1, alcohol 10). Lepra-bacilli stain; tubercle-bacilli do not.

c. Trichomycetes.

1. Actinomyces. Alcohol and formol fixation are best. Good preparations can be obtained with hæmatoxylin and eosin, Van Gieson’s or Weigert-Gram’s. The special staining methods advised give no better results than these simpler stains. Differential staining of clubs and mycelium may be obtained by Mallory’s method:—

1. Stain lightly in alum-cochineal. (Powdered cochineal 6 grms., ammonia alum 6 grms., water 100 cc. Boil half an hour, add water lost by evaporation, filter, add crystals of thymol.)

2. Saturated watery eosin 10 minutes.

3. Wash in water.

4. Stain in aniline gentian-violet 2-5 minutes.

5. Wash in physiologic saline solution.

6. Transfer sections to Lugol’s for 1 minute.

7. Pass rapidly through water.

8. Dry thoroughly between folds of filter-paper.

9. Cover section with aniline-oil until clear.

10. Xylol; balsam.

Clubs pink; mycelium blue.

2. Nocardia, Cladothrix, Streptothrix and Leptothrix. Löffler’s methylene-blue and carbol-fuchsin give good results. The Nocardiæ are acid-fast with dilute acids. They give good preparations with Weigert’s fibrin stain and lithium-carmine.

d. Vibrios.

1. Cholera Vibrios. Gram-negative. Sections may be stained with fuchsin or methylene-blue.

e. Spirilla and Spirochætes.

1. Spirillum of Recurrent Fever. Stain in sections with Levaditi’s silver-method, or with Nikiforoff’s method:—

1. Fix for 24 hours in equal parts of a 5 per cent water solution of potassium bichromate and a saturated solution of mercuric chloride in 0.6 per cent sodium chloride.

2. After-harden in graded alcohols in the incubator.

3. Imbed in paraffin.

4. Stain 24 hours in a mixture of alcoholic 1 per cent solution of tropæolin 5 cc., concentrated watery methylene-blue solution 10 cc., caustic potash solution (1:1000) 2 drops.

5. Wash in water.

6. Dip several times in a mixture of equal parts of absolute alcohol and ether.

7. Oil of bergamot; xylol; balsam.

The spirillum of African relapsing fever stains with the same stains as the spirillum Obermeieri. The spirochætes of Vincent’s angina and fowl-spirillosis, and the spirochæte refringens stain with watery aniline dyes and with Giemsa’s stain; in section they are stained by the Levaditi method.

2. Spirochaeta Pallida (Treponema Pallidum). This organism is best examined in the living condition by means of the dark-field illumination (dark-field condenser). A very simple method of dark-field illumination consists of the use of India ink. The suspected discharge or serum is placed on a slide and an equal quantity of ink (Gunther’s or Higgin’s) added. The serum and ink are rapidly mixed and spread over the slide to dry in a pale brown smear. The oil for the immersion is placed directly on the smear. The spirochætes appear as white spirals against a brownish-black field. The best results are obtained with serum; the presence of mucus or fibrin interferes with the clearness of the picture obtained.

Smears of serum from syphilitic lesions may be dried in the air and fixed in absolute alcohol or equal parts of absolute alcohol and ether for 15-20 minutes. They may then be stained by Giemsa’s (old formula) stain (azur II-eosin 3 grms., azur II 0.8 grm., glycerin [Merck’s chemically pure] 250 grms., methyl-alcohol [Kahlbaum I] 250 grms.). This solution can be obtained from Grübler. Ten drops of the stain are mixed with 10 cc. of distilled water immediately before the staining. The fixed preparation is covered with the diluted staining fluid and warmed over the flame until a slight steam arises. It is then allowed to cool for about 15 seconds, when the stain is poured off and replaced by fresh, and the process repeated four or five times, when the preparation is washed, dried and mounted in balsam. Spirochætes are dark red. Slide or cover-glass and forceps must be absolutely clean. Smears may also be fixed and stained by Wright’s blood-stain.

For the demonstration of the treponema in sections the method of Levaditi gives the most satisfactory results:—

1. Fix thin pieces of tissue 24 hours or longer in 10 per cent formol. (Formol-Müller’s and alcohol-fixation may also be used.)

2. 24 hours in 96 per cent alcohol.

3. Transfer to distilled water until tissue sinks.

4. Impregnation for 3 days in incubator, in a 1.5-3 per cent silver nitrate solution.

5. Wash for a short time in water.

6. Reduce for 48 hours, in the dark, at room-temperature, in pyrogallic acid 4 grms., 40 per cent formol 5 cc., distilled water 100 cc.

7. Wash in water. Cut on freezing-microtome, or imbed in celloidin or paraffin. Toluidin-blue or safranin may be used as a contrast-stain.

The spirochætes are dark brown to black. Silver precipitates occur chiefly in the outer portions of the tissue. The reticulum is brown; other parts of the tissue are yellowish. Levaditi’s more recent modification of this method does not give so good results as the original.

Schmorl’s Staining of Sections with Giemsa’s Stain.

1. Fix in 10 per cent formol. Cut very thin sections on freezing-microtome.

2. Place the sections in a staining dish containing a measured amount of distilled water. To each cc. of water add one drop of Giemsa’s stain. Use clean glass-needles to manipulate the sections. After 1 hour transfer sections to a fresh solution, in which they are left 5-12-24 hours.

3. Wash quickly in a concentrated solution of potassium alum, then quickly in water.

4. Mount in glycerin-gelatin; or dry on the slide until nearly perfectly dry, then xylol, and balsam, or cedar oil. Alcohol must not be used.

II. THE STAINING OF PATHOGENIC YEASTS AND MOULDS IN SECTIONS.

1. Blastomycetes. The parasites of blastomycetic dermatitis can be demonstrated unstained in pus treated with a weak sodium hydroxide. In sections they are easily found after treatment with ordinary staining methods. The various modifications of the Romanowsky method, or other methylene-blue-eosin staining, give better staining of the parasite than can be obtained by hæmatoxylin and eosin.

2. Oïdium Albicans. Staining with Weigert-Gram’s and lithium-carmine gives beautiful preparations.

3. Moulds. These are best examined in the unstained condition, by treating the material with equal parts of alcohol and ether, followed by a 3 per cent potassium hydroxide solution. The organisms and spores are brought out distinctly. Löffler’s methylene-blue may be used for staining. In the case of sections stain 1-2 hours and contrast with eosin. For the examination of hairs or horny scales for fungi, Unna’s method may be used:—

1. Add glacial acetic acid to hair or epidermis; make cover-glass preparations, drying by heat.

2. Ether and alcohol equal parts.

3. Stain in borax 1 grm., methylene-blue 1 grm., water 100 cc., ½-5 minutes.

4. Wash in water; dry; balsam.

If the horny elements are too deeply stained, decolorize in 1 per cent acetic for 10 seconds, or in 1 per cent oxalic, citric, or arsenious acid for 1 minute.

III. THE STAINING OF ANIMAL PARASITES.

1. Amoeba Coli. Examine fresh material from fæces, abscesses or cultures, in physiologic saline solution, on a warm stage. Stain under the cover with methylene-blue and carmine. Make permanent mounts by removing excess of stain and running in 50 per cent glycerin. In fixed preparations the nuclei of the amoebæ do not stain with ordinary nuclear stains. Mallory’s method may be used:—

1. Fix in alcohol.

2. Stain sections in a saturated aqueous solution of thionin 3-5 minutes.

3. Differentiate in a 2 per cent aqueous solution of oxalic acid for ½-1 minute.

4. Wash in water; dehydrate in absolute alcohol; clear in xylol; mount in xylol-balsam.

Nuclei of the amœbæ and granules of the mast-cells are brownish-red; nuclei of cells blue.

2. Trichomonas vaginalis and intestinalis; Cercomonas coli; Megastoma entericum; Balantidium coli; Pyrosoma bigeminum; Trypanosoma; Leishman-Donovan bodies, and allied forms are best stained with the modifications of Romanowsky’s stain; the chromatin is red-violet (macro-nucleus red, micro-nucleus black, flagellum red, protoplasm blue, basophilic granules black). For staining in sections mercuric chloride or Zenker’s fixation followed by staining with polychrome methylene-blue, Giemsa’s or the modifications of the Romanowsky method may be employed.

3. Plasmodium Malariae. For films make medium smears (not too thin); fix with equal parts of absolute alcohol and ether for ½-1 hour; or fix and stain in the same solution (Leishman-Romanowsky, Wright’s stain, etc.). For single staining methylene-blue, carbol-thionin, etc., may be employed; for double staining eosin and methylene-blue. Ehrlich’s tri-acid, or any of the eosin-methylene-blue combinations may be used (particularly the Leishman-Romanowsky or Wright’s). With the Romanowsky methods the body of malarial organism is stained blue, the chromatin varying shades of lilac, red, purplish-red or almost black. When the blood contains but few parasites 1 cc. may be drawn, mixed with 20 cc. of distilled water and centrifugated. Smears are then made of the sediment. For the staining of the plasmodium in imbedded tissues the following method is recommended by Bignami. The tissue should be fixed in formol or mercuric chloride, preferably a mixture of mercuric chloride 1 grm., sodium chloride 0.75 grm., acetic acid 0.75 grm., water 200 cc. Fix for 2 hours; after-harden in alcohol and iodine-alcohol, changing the alcohol each day for seven days. Dehydrate in absolute alcohol, and imbed in celloidin or paraffin. Stain in a saturated watery solution of magenta or in a mixture of equal parts of saturated alcoholic mixtures of magenta and orange G. Good results may, however, be obtained with Löffler’s methylene-blue. Clear in xylol; mount in balsam.

4. Coccidia and Sarcosporidia. The ordinary fixations give good results. Imbed in paraffin or celloidin. Weigert’s iron-hæmatoxylin and Van Gieson’s give as good pictures as any of the special methods advised.

5. Negri Bodies of Rabies. Examine in smears or make sections. Take portions of gray brain-substance from the cortex in the region of the fissure of Rolando (in the dog from around the crucial sulcus), from the hippocampus, and from the cerebellum. Smear cover or slide by taking a thin slice of the gray matter and compressing it between two slides, or cover and slide, or by drawing the cover across the cut surface in order to get some of the cells. Dry in the air; stain with watery methylene-blue; wash; stain with watery acid fuchsin; wash in water; blot dry; mount in balsam. Negri bodies fuchsin-red (about size of red blood cells); everything else blue. When dried in the air and then fixed in methyl alcohol for 5 minutes the smears may be stained by Giemsa’s method. For the demonstration of the bodies in sections fix in Zenker’s, imbed in paraffin, and stain by the eosin-methylene-blue method. The bodies take the eosin stain. Formol fixation, freezing-microtome and Romanowsky stain give quick results. Mann’s method for the staining of Negri bodies in sections is strongly recommended by many workers. Fix material in mercuric chloride or Mann’s fluid (1 grm. picric acid and 2 grms. tannin dissolved in 100 cc. concentrated water solution of mercuric chloride) for 24 hours; wash thoroughly in running water; imbed in paraffin. Stain in Mann’s mixture (1 per cent aqueous methyl-blue [not methylene-blue] 35 parts, 1 per cent aqueous eosin 35 parts) for 24 hours; wash in water; rinse in absolute alcohol; place in alkaline alcohol (absolute alcohol 50 cc., 4 drops of a 1 per cent solution of sodium hydroxide) for 15-20 seconds until sections become reddish; wash quickly in alcohol; wash about 2 minutes in water until superfluous color is removed; place in weak acidulated water (acetic acid) 1-2 minutes until sections are blue; quick dehydration in alcohol; xylol; balsam. Cells are blue, nucleoli and blood-vessels red; Negri bodies bright red. For quick diagnosis use acetone fixation and imbedding, stain in Mann’s fluid 2-4 minutes, and proceed as in the Mann’s method. While these bodies possess a great diagnostic importance for rabies, their exact nature must still be regarded as unsettled; they are most probably not parasites.

6. Vaccine Bodies. Fix in Flemming’s, mercuric chloride or Zenker’s; imbed in celloidin or paraffin. Stain with Heidenhain’s iron-hæmatoxylin (bodies black) or Biondi-Heidenhain mixture (bodies blue, nuclei of leukocytes and mitoses green, nuclei of epithelium and connective-tissue blue, protoplasm and connective-tissue red). These bodies are probably not parasites, but may be products of cell-degeneration.

7. Vermes. The heads, proglottides and ova are best examined in the fresh state, in physiologic saline or glycerin. Acetic acid may be used to bring out details. Berlin-blue or methylene-blue may be injected through the genital pore for the demonstration of the excretory and genital organs. Scolices and hooklets of echinococcus may be obtained by scraping the cyst-wall; examine in glycerin. Permanent preparations of cestodes, nematodes and trematodes may be made by fixing in mercuric chloride, formol or Flemming’s, after-hardening in alcohol, staining in orange G, borax carmine, alum hæmatoxylin, hæmatoxylin and eosin etc., mounting in glycerin gelatin; or dehydrating, clearing in xylol and mounting in balsam. For sections imbed in paraffin or celloidin. Trichinæ may be studied by teasing the fresh muscle; by digesting with pepsin and hydrochloric acid and examining the freed trichinæ on a warm stage; or by imbedding in paraffin or celloidin and staining with hæmatoxylin and eosin. Permanent mounts of the embryos of filaria may be made by fixing cover-glass preparations of blood or chylous fluid by heat or mercuric chloride, and staining for a few seconds with Löffler’s or a 2 per cent aqueous thionin.

CHAPTER XXVIII.
THE STAINING OF SPECIAL ORGANS AND TISSUES.

I. BLOOD AND BLOOD-FORMING ORGANS.

The blood may be examined by means of films, stained or unstained, or by sections, celloidin or paraffin.

A. FILMS. The blood may be obtained from the pulp of the ring finger, from the skin over the knuckles, or from the posterior aspect of the lobe of the ear. The place selected should be carefully cleansed with water, soap and 1/1000 mercuric chloride solution, and finally with alcohol and ether. A puncture is made with a sterilized triangular needle or knife, or a stub-pen with one point broken off. The last-named makes a most useful and inexpensive instrument for this purpose. The puncture should be made by a quick and deep stab, so that sufficient blood can be obtained from one stab-wound. Pressure should not be employed to force blood from the wound. Bleeding may be encouraged by letting the arm hang down, or by applying pressure in the furrow of the terminal joint of the finger. The first drop of blood should be wiped away with a clean towel. When the second drop reaches the size of a pin-head touch it with the under side of a perfectly clean cover-glass, held by forceps, not by the fingers; place this cover-glass immediately upon another clean cover, so that the blood will spread out between the two covers in a thin film. The covers are then separated by sliding them apart without pressing or squeezing; place covers with film side upward, and dry in the air. The films should not be touched with the hands; forceps alone should be used to handle them. If the blood does not dry as quickly as it is spread the film will be too thick. Films may be made upon slides in the same way, or the drop of blood may be caught upon the edge of a clean cover, slide or “spreader” and then drawn rapidly across a slide. The dried film may be marked by scratching with a needle-point the number and date on the film itself. Blood-films may be fixed without drying by exposure to the vapor of formol or osmic acid for several seconds and then dropping into absolute alcohol. Formol alcohol, saturated mercuric chloride solution or Flemming’s solution may also be used for the fixation of wet films, fixing for 5-10 minutes, and washing thoroughly after each of the last two solutions. The dried film may be fixed by exposure to heat (110-115°C.) for 5-10 minutes for Ehrlich’s triple stain, and for 2 hours for the methylene-blue-eosin methods; 30-60 seconds at a temperature of 120°C. may suffice; the film should be brought at once into the required temperature. Heat-fixed films are improved by dipping them for a few minutes in mercuric chloride solution and then washing well before staining. Acetone-free methyl alcohol (1-2 minutes), absolute alcohol and ether in equal parts (½-12 hours), formol-alcohol (1-2 minutes), alcoholic mercuric chloride (absolute alcohol 25 cc., ether 25 cc., 5 drops of a 2 grms. mercuric chloride solution in 10 cc. of absolute alcohol) for 2-5 minutes, and formol-vapor are the chief solutions used for fixing the air-dried film. For ordinary work methyl alcohol, formol alcohol, and the absolute-alcohol and ether mixture give good results; heat fixation brings out the granules well, and mercuric chloride is a good fixative for the leukocytes. The combination of fixation and staining, as in Leishman’s or Wright’s modification of the Romanowsky method, is also recommended for general work.

For the staining of blood-films an almost endless variety of staining-methods can be found in the literature. Many of these represent slight deviations in the method of making the stain or in its application, such deviations marking stages of improvement in the development of the method. It is not necessary, therefore, to give all of these methods, but to consider only the latest modifications of value. In a general way blood-stains may be divided into five classes:—

1. HAEMATOXYLIN AND EOSIN.

Fix in equal parts of absolute alcohol and ether for at least 30 minutes; stain with hæmalum and eosin, or Ehrlich’s acid hæmatoxylin and eosin. By adding O.5 grm. of eosin to the formula for Ehrlich’s acid hæmatoxylin a combination stain can be made that is very good for blood-films fixed by heat or absolute alcohol and ether. Stain 2-24 hours, wash, dry and mount in xylol balsam.

2. EOSIN AND METHYLENE-BLUE.

Fix by formol (dried film over 40 per cent formol for 1 minute); absolute alcohol for 1 minute; stain 5 minutes in a 1 per cent watery eosin; then without removing eosin place in watery methylene-blue for 2 minutes; wash quickly; dry in air; balsam.

3. MIXTURES OF EOSIN AND METHYLENE-BLUE.

The numerous mixtures of methylene-blue and eosin are not very stable, can be kept for a few days only, and give varying results. Jenner improved this method of staining greatly by collecting the precipitate formed by the addition of eosin to methylene-blue, and dissolving it in pure methyl alcohol, thus giving a solution that fixes and stains at the same time. The May-Grünwald method is practically the same.

Jenner’s Method.

a. Water-soluble eosin, 1.25 grms.
Distilled water, 100 cc.
b. Medicinal methylene-blue, 1 grm.
Distilled water, 100 cc.

Mix equal parts of a and b in an open basin, stirring with a glass rod. Let stand for 24 hours; filter; dry the residue at 50°C. Wash residue thoroughly with distilled water and again dry thoroughly. Take 0.5 grm. of the dried powder and dissolve in 100 cc. of pure methyl alcohol. Filter. Solution keeps well.

1. Make blood-film. Dry in air. Do not fix.

2. Cover film with stain, keeping under watch-glass to prevent evaporation Stain 2 minutes.

3. Wash in distilled water until the film has a pink color. Dry in air. Mount in xylol-balsam.

Neutrophile granules are red, eosinophile rose red, basophile granules violet, red blood cells and central portion of blood-platelets are terra-cotta, leukocyte nuclei and granules in red blood cells are blue, protoplasm of nuclei and outer portion of platelets light blue.

4. MODIFICATIONS OF THE ROMANOWSKY METHOD.

A large group of stains has resulted from various applications of the Romanowsky idea of uniting equimolecular proportions of methylene-blue and eosin, and the solution of the dyes so obtained in some suitable solvent. These dyes consist of mixtures of methylene violet, methylene azure, eosinate of methylene blue, etc., and can be obtained from Grübler and Co. under various names, such as Azur-blau, Bleu Borrel, Giemsa’s stain, Leishman’s stain, etc. Hastings, Leishman, Wright and others have combined the Romanowsky method with that of Jenner by dissolving the new dyes obtained by their various modifications in pure methyl alcohol, so as to form a solution that will fix and stain at the same time. Hastings’ stain is a modification of Nocht’s stain; Wright’s stain is a modification of the Leishman-Romanowsky method. The revised directions given by Wright for making and using his stain are here given. Wright’s method and the Giemsa stain possess all of the staining advantages afforded by the variations of the Romanowsky method, and are alone given here. The former is recommended for blood-work, the latter for the staining of protozoa.

Wright’s Blood-stain.

To a 0.5 per cent aqueous solution of sodium bicarbonate add methylene-blue (B.X or medicinal) in the proportion of 1 grm. of the dye to each 100 cc. of the solution. Heat the mixture in a steam sterilizer at 100°C. for one full hour, counting the time after the sterilizer has become thoroughly heated. The mixture should be placed in a flask of such size and shape that the fluid will not be more than 6 cm. deep. After heating, allow the mixture to cool, placing the flask in cold water if desired, and then filter it to remove the precipitate. When cold the fluid should have a deep purple-red color when viewed in a thin layer by transmitted yellowish artificial light. It does not show this color while warm.

To each 100 cc. of the filtered mixture add 500 cc. of a 0.1 per cent aqueous solution of yellow water-soluble eosin, and mix thoroughly. Collect on a filter the abundant precipitate which immediately appears. When the precipitate is dry, dissolve it in pure methyl alcohol (Merck’s) in the proportion of 0.1 grm. to 60 cc. of the alcohol. To facilitate solution the precipitate is to be rubbed up in a porcelain dish or mortar with a spatula or pestle. This alcoholic solution is the staining solution. It should be kept in a tightly-stoppered bottle. Should it become concentrated through evaporation methyl alcohol in proper quantity should be added.

1. Cover film with a given quantity of staining fluid by means of a medicine dropper.

2. After 1 minute add to the staining fluid on the film the same quantity of distilled water by means of the medicine dropper, and allow the mixture to remain for 2-3 minutes according to the intensity of the stain desired. A longer period of staining may produce a precipitate. Eosinophile granules show best after short staining. The quantity of diluted stain on the preparation should not be so great that some of it runs off.

3. Wash the preparation in water for 30 seconds or until the thinner portions of the film become yellow or pink in color.

4. Dry, and mount in balsam.

Films more than a few hours old do not stain as well as fresh ones.

The red cells are orange or pink in color. Polychromatophilia and punctate basophilia or granular degeneration are well shown. Nucleated reds have deep-blue nuclei, and their cytoplasm is usually bluish. The lymphocytes have dark purplish-blue nuclei and cytoplasm of a robin’s-egg blue, in which a few dark-blue or purplish granules are sometimes present. The nuclei of the polynuclear neutrophilic leukocytes are dark-blue or dark lilac-colored, the granules reddish-lilac. The eosinophiles have blue or dark lilac nuclei, a blue cytoplasm and eosin-red granules. The large mononuclear leukocytes have a dark lilac or blue nucleus, cytoplasm pale blue or blue with dark-lilac or deep purple granules. Mast-cells have purplish or dark-blue nuclei, bluish protoplasm and coarse dark purple or black granules. Myelocytes have dark blue or lilac nuclei, blue cytoplasm, and dark-lilac or reddish-lilac granules. Blood platelets are blue with small violet or purplish granules in their central portions. Malarial parasites have a blue body and lilac or red chromatin. Spirochæte pallida is pale blue.

Giemsa’s Method.

a, One per cent water solution of azur-blau; b, one per cent watery solution of eosin. For staining take 1 cc. of b, add 10 cc. of water, and then 1 cc. of the azur-blau solution. Stain 10 minutes to 1 hour.

Giemsa’s Old Method.

Azur II—Eosin3.0grm.
Azur II0.8grm.
Glycerin (Merck’s pure)250.0cc.
Methyl-alcohol (Kahlbaum I)250.0cc.

To 1 cc. of distilled water in a small, perfectly clean graduate add 1 drop of the stain, shaking very gently. Make very thin film; dry in air; fix 15-20 minutes in absolute alcohol. Cover preparation with a thin layer of the freshly diluted stain for 10-15 minutes, renewing stain at end of 10 minutes. Wash in a stream of water. Differentiate over-stained preparations in distilled water. Dry with absorbent paper; mount in balsam. Stains the spirochæte pallida and malarial organisms. The Giemsa solution may be obtained from Grübler. A more intense staining can be obtained by adding to the water used for diluting the stain 1-2 drops of a 0.1 per cent solution of potassium carbonate.

5. SPECIAL ELECTIVE STAINS FOR THE BLOOD.

1. Ehrlich’s Triple Stain.

Saturated watery solution of Orange G120cc.
Saturated watery solution of acid fuchsin80cc.
Saturated watery solution of methyl green100cc.
Glycerin50cc.
Distilled water300cc.
Absolute alcohol180cc.

Mix gradually; allow to stand for several months; do not shake or filter. Remove stain with pipette. Fix by heat, or pure methyl alcohol for 5 minutes. Stain 5-10 minutes; wash thoroughly, dry and mount in balsam. Neutrophile granules violet; eosinophile, a bright red; nuclei of the neutrophilic and eosinophilic cells greenish-blue; nuclei of the lymphocytes deep-blue; nuclei of the large mononuclears pale blue; those of red cells intense blue; red cells copper red. The Aronson-Philipp modification is more variable and less satisfactory.

Pappenheim’s Stain for Lymphocytes.

3-4 parts of polychrome methylene-blue or methyl green to 1-2 parts of pyronin. Fix in absolute alcohol. Nuclei blue-green; protoplasm bright red.

Staining of Blood-platelets.

The blood-platelets may be examined in the fresh state by coating a cover-slip with Deetjen’s agar-solution (boil 5 grms. agar-agar in 500 cc. distilled water, filter hot, and to each 100 cc. of the filtrate add 0.6 grm. sodium chloride, 6-8 cc. of a 10 per cent solution of sodium phosphate and 5 cc. of a 10 per cent solution of sodium diphosphate). Place drop of blood on this coating and examine on warm stage. For permanent stained preparations bleed into a fixing and staining fluid (equal parts alcohol and ether and Romanowsky’s stain) or use Wright’s stain.

Bremer’s Diabetic Reaction.

Take a clean cover-glass, smear one-half with normal blood, the other half with diabetic blood. Fix for 2 hours at 120°C., or in equal parts of absolute alcohol and ether at 60°C. for 4 minutes. Stain in a 10 per cent watery methylene-blue for 2 minutes, wash off the stain in water, and stain for 10 seconds in a ⅛ per cent watery eosin. Wash, dry and mount in balsam. In diabetic blood the red cells are green; in normal blood red. While this reaction is constant in diabetic blood it also occurs in leukæmia, Hodgkin’s disease, exophthalmic goitre and multiple neuritis. A 1 per cent solution of Biebrich scarlet stains diabetic blood intensely, normal blood but slightly. On the other hand, a 1 per cent methylene-blue and a 1 per cent Congo red stain normal blood intensely and diabetic blood slightly.

Staining of Glycogen in Leukocytes.

To a solution of Lugol’s (100:3:1) add sufficient gum arabic to make a syrupy mixture. Keep tightly corked. Place a drop of this solution upon an air-dried film. After 1 minute dry with blotting paper. Examine with oil immersion. A positive reaction is shown by the presence of a diffuse brown or reddish-brown coloration or granules in the cell-body of the polymorphonuclear leukocytes.

Staining of Fat in Blood.

Stain in solutions of scharlach R or sudan III in 70 per cent alcohol.

Staining of Blood-parasites.

The malarial parasites, trypanosomes, Leishman-Donovan bodies, sporidia, piroplasma bigeminum, spirilla and spirochætes and the filaria may all be stained with Wright’s or Giemsa’s modification of the Romanowsky method, or by any of the modifications of this method. (See also Staining of Animal Parasites.)

B. SECTIONS. The blood is allowed to drop directly into Flemming’s solution and allowed to stand for 24 hours. It is then washed in water by repeated decanting, or the coagulum may be placed in a bottle covered with muslin, and then exposed to running water. It is after-hardened in alcohol and imbedded in paraffin. Safranin should be used to stain the sections. This method is especially good for the demonstration of mitoses in the blood-cells.

Bone-marrow.

Prepare films and fix and stain, as for blood films. For sections, fix the marrow in formol-Müller’s, mercuric chloride, Zenker’s, etc.; imbed in paraffin; cut very thin sections; stain with Ehrlich’s triple stain or Wright’s modification of Leishman’s stain. To distinguish the young forms of erythrocytes and leukocytes Trambusi fixes in Flemming’s, stains the sections in a 1 per cent thionin solution in aniline-water (4:100), differentiates in acid-alcohol, and then brings the sections into a watery eosin and finally an alcoholic eosin, and mounts in xylol-balsam.

Spleen and Lymphnodes.

Fresh material may be obtained by means of a trocar, and may be examined in the fresh state, or films may be prepared, fixed and stained, as for blood-smears. Sections of fixed tissues may be obtained by the use of the same fixing and staining methods employed in the study of the blood or bone-marrow. For the study of the reticulum Mallory’s reticulum stain or the digestion-method may be used. In ordinary work formol-fixation followed by eosin-staining is of great value in distinguishing hæmolymphnodes and lymphatic glands.

II. BONE.

For ordinary work decalcification is necessary except for those pathologic conditions in which the lime-salts have been lost (see Chapter [XXI]). Imbed in celloidin preferably. When decalcification has been carried out place sections in an alkaline solution before staining, and stain for a longer period than usual. Methylene-blue and eosin stain osteoblasts and osteoclasts very well. Van Gieson’s is an especially useful stain for ordinary work; osteoid tissue is red, calcified areas yellow. Sections of bone without decalcification can be prepared by fixing, hardening and staining in bulk; the bone is then sawn in the dried condition, and the sections ground down to the required thickness. Schmorl’s methods for the preparation of bone-sections have practically superseded all other staining methods.

Schmorl’s Thionin-picric-acid Method.

1. Fix in formol or formol-Müller’s preferably.

2. Decalcify in formol-nitric acid or Ebner’s alcoholic HCl acid solution.

3. Wash thoroughly in water. After-harden in increasing strengths of alcohol; freeze or imbed in celloidin (not paraffin); cut.

4. Transfer sections to water for 10 minutes.

5. Stain sections, well spread out, for 5-10 minutes in a solution of 2 cc. of a saturated solution of thionin in 50 per cent alcohol and 10 cc. of water, to which 1-2 drops of ammonia are added.

6. Wash in water.

7. Stain ½-1 minute in hot saturated and cold filtered watery-solution of picric acid.

8. Wash in water.

9. Differentiate in 70 per cent alcohol until the color ceases to come away in blue-green clouds, 5-10 minutes or more.

10. Dehydrate in 96 per cent alcohol.

11. Clear in phenol xylol; xylol; balsam.

Lacunæ and canaliculi dark brown to black; bone cells red; ground substance yellow or brownish yellow. Calcified areas take a darker yellow than non-calcified. This method consists in an impregnation with a fine precipitate rather than a staining. Should the precipitate be too heavy in portions of the section it may be removed by thorough washing between 9 and 10.

Schmorl’s Thionin and Phosphotungstic or Phosphomolybdic Acid Method.

1. Fix thin pieces of fresh bone in formol, then in Müller’s for 6-8 weeks, or 3-4 weeks in the incubator. Fixation is best at 37°C.

2. Decalcify in Ebner’s hydrochloric acid solution. Wash thoroughly. After-harden in alcohol. Imbed in celloidin or paraffin. Cut very thin sections.

3. Water for 10 minutes.

4. Stain in the alkaline-thionin solution, as in previous method, for 3 minutes.

5. Wash in water.

6. With glass needles transfer sections to a saturated watery solution of phosphotungstic or phosphomolybdic acid for a few seconds.

7. Wash 5-10 minutes, until section is sky-blue in color.

8. Fix the stain for 3-5 minutes in ammonia 1 part, water 10 parts.

9. Transfer directly to 90 per cent alcohol; change twice.

10. Dehydrate; clear in carbol-xylol; balsam.

If the ground-substance is too dark differentiate in acid alcohol before dehydrating, and then wash thoroughly before beginning the dehydration.

Outlines of lacunæ and canaliculi are dark blue; ground-substance a light or greenish blue; cellular elements a diffuse blue. Schmorl advises this method for growing bone; in rickets the well-ossified areas alone stain well. Both of the Schmorl methods can be used for the study of teeth as well.

Staining of Sharpey’s Fibres (v. Kölliker).

1. Harden, decalcify, imbed, cut.

2. Place section in strong acetic acid until it becomes transparent.

3. Stain in a saturated watery solution of indigo carmine for 15-16 seconds.

4. Wash in water; mount in glycerin.

Fibres red; ground-substance blue.

III. CARTILAGE.

Cartilage stains deeply with hæmatoxylin; with Weigert’s fibrin stain it holds the blue; with thionin and polychrome methylene-blue stains for mucin cartilage stains metachromatically red.

IV. CONNECTIVE TISSUES.

a. Connective-tissue Fibrils. The demonstration of connective-tissue fibrillæ or reticulum is of great importance in the differential diagnosis of sarcoma and carcinoma. Van Gieson’s method is the best stain for the coarser fibrils, but does not bring out the finer reticulum as well as Mallory’s aniline-blue method.

1. Mallory’s Reticulum Stain.

1. Fix in mercuric chloride or Zenker’s. After-harden in alcohol; imbed in celloidin or paraffin; cut.

2. Stain in 1/10 aqueous acid fuchsin 5-10 minutes.

3. Transfer directly to the following solution and stain for 20 minutes or longer:—

Aniline-blue, water-soluble (Grübler)0.5grm.
Orange G (Grübler)2.0grms.
1 per cent aqueous solution of phosphomolybdic acid100cc.

4. Wash and dehydrate in several changes of 95 per cent alcohol; dry with absorbent paper.

5. Clear in xylol or origanum oil.

6. Mount in balsam.

Fibrils and reticulum of connective tissue, amyloid, mucin and connective-tissue hyalin are blue; nuclei, protoplasm, fibroglia fibres, axis cylinders, neuroglia fibres and fibrin are red; elastic fibres are pink or yellow; red blood-cells and myelin-sheaths are yellow.

2. Mall’s Method for Reticulum.

1. Digest frozen sections of fresh tissue for 24 hours in a solution of Parke, Davis and Co.’s pancreatin 5 grms., soda bicarbonate 10 grms., water 100 cc.

2. Wash carefully in water.

3. Place section in test-tube half-full of water and shake thoroughly to remove cells.

4. Spread out on slide and allow to dry.

5. Allow a few drops of the following solution to dry on slide:—picric acid 10 grms., absolute alcohol 33 cc., water 300 cc.

6. Stain for 30 minutes in the following mixture: acid fuchsin 10 grms., absolute alcohol 33 cc., water 66 cc.

7. Wash in the picric acid solution for a second.

8. Dehydrate in absolute alcohol; xylol; balsam.

3. Unna’s Method for Collagen.

1. Harden in absolute alcohol; imbed; cut.

2. Stain 5-15 minutes in polychrome methylene-blue.

3. Wash in water.

4. Differentiate in 1 per cent neutral orcein in absolute alcohol, 15 minutes.

5. Dehydrate in absolute alcohol.

6. Clear in xylol; mount in balsam.

Nuclei dark blue; protoplasm light blue; collagen dark red; plasma-cell granules greenish-blue; mast-cell granules red.

4. Mallory’s Fibroglia Method.

1. Fix thin, small, fresh pieces of tissue in Zenker’s fluid; harden in alcohol; imbed in celloidin or paraffin; cut.

2. Stain sections in 1 per cent aqueous acid fuchsin for 12 hours in the cold, or 20-30 minutes at 50-56°C.

3. Wash in water for 5 seconds.

4. Differentiate in 0.25 per cent aqueous potassium permanganate solution 10-20 seconds.

5. Wash in water for 5 seconds.

6. Dehydrate in alcohol; clear in xylol or origanum; mount in balsam.

Fibroglia fibrils and cell-nuclei intensely red; contractile elements of striped muscle, smooth muscle, neuroglia fibres, cuticular surfaces of epithelial cells and fibrin are also red; connective-tissue fibres brownish-yellow or colorless; elastic fibres, unless degenerated, bright yellow.

b. Elastic Fibres. Weigert’s method is so superior to the Unna orcein-stain that it alone is given here. It is our best elective stain: gives permanent preparations, and is in every way practical. The stain keeps well.

Weigert’s Method for Staining Elastic Fibres.

Preparation. Boil in a porcelain dish resorcin 4 grms., fuchsin (Grübler) 2 grms., and water 200 cc. After the mixture has boiled a few seconds add 25 cc. of liquor-ferri sesquichlor., Pharm. Germ. III. Stir well and boil for 5 minutes. When cool, filter. Carefully loosen the filter from the funnel, transfer it to the same porcelain dish which still contains a small amount of sediment, and add 200 cc. of 94 per cent alcohol. Boil and stir carefully. Remove the filter-paper when all the sediment is dissolved. Cool, filter; make up the filtrate to 200 cc. with 94 per cent alcohol, and to these 200 cc. add 8 cc. of hydrochloric acid. Resorcin-fuchsin may be obtained from Grübler, but the freshly-prepared stain gives better results.

1. Fix in any ordinary solution; imbed; cut.

2. Stain with lithium-carmine and differentiate in acid alcohol; wash thoroughly.

3. Stain in the resorcin-fuchsin mixture for 20-60 minutes.

4. Wash rapidly in acid alcohol.

5. Dehydrate and differentiate in absolute alcohol until section is red.

6. Clear in xylol; balsam.

Nuclei are red; elastic fibres blue-black. Should the stain when old give a diffuse staining differentiate for a longer time in acid alcohol.

c. Fat Tissue. Use same methods as advised for the demonstration of fatty degeneration and infiltration (osmic acid, scharlach R, sudan III).

V. EAR.

Remove temporal bone; fix in formol-Müller’s; decalcify in trichloracetic acid; wash; after-harden in alcohol; imbed in celloidin. For nerve-endings use Golgi’s and methylene-blue methods.

VI. EYE.

Fix in Müller’s, formol-Müller’s, Zenker’s, formol, Flemming’s or Marchi’s solution. Aid fixation by incisions into sclera. The eye should not be left in formol for more than 3 days. Section as desired; imbed larger pieces in celloidin, small ones in paraffin. Use alum-carmine, iron-hæmatoxylin, Van Gieson’s, Weigert’s elastic stain, Levaditi’s silver-method, Golgi’s nerve-stain, methylene-blue method, etc., according to the object of the investigation.

VII. LIVER.

For the demonstration of the bile-capillaries Weigert’s neuroglia method gives the best results. (See Page [300].) This method may be used with sections cut on the freezing-microtome after fixation in formol. Such sections are placed in a ½ per cent solution of chromic acid for 1 hour, transferred to the neuroglia mordant for 5-6 hours, washed well with water, and then treated as for the neuroglia method. Van Gieson’s method may also be used for frozen sections of formol-fixed tissue. The walls of the capillaries show as fuchsin-colored streaks.

VIII. MUSCLE.

Van Gieson’s is the best general stain for both striped and unstriped muscle, as it differentiates the muscle perfectly from the connective-tissue. Mallory’s reticulum stain may also be used for the same purpose. For the study of myoglia fibrils the tissue must be fixed within a few minutes after its removal from the living body. Autopsy material cannot be used. These fibrils can be demonstrated by Mallory’s fibroglia stain, or by Mallory’s phosphotungstic-acid hæmatoxylin stain for neuroglia. (See below.)

IX. NERVOUS SYSTEM.

It is impossible in a book on general pathologic technic to consider all of the numerous staining methods that have been devised for the study of the nervous system. I have attempted, therefore, to pick out the best selective methods for the staining of the more important nervous structures, so as to cover adequately the general held of nervous pathology. Formol has now replaced Müller’s for the preliminary fixation of nervous tissue, because of its quick action, and because after its use chromic acid may be employed to mordant the tissue, when it is desired to use certain staining methods requiring such mordanting. Celloidin imbedding is preferable, although paraffin may be used for general work. General stains, such as hæmatoxylin and eosin, and Van Gieson’s are used for general impressions.

1. METHODS FOR STAINING GANGLION CELLS.

A. Lenhossék’s Method.

1. Fix in equal parts of saturated watery picric acid and mercuric chloride (Rabl’s mixture); after-harden in absolute alcohol; imbed in paraffin; cut.

2. Stain in saturated watery solution of toluidin blue, or thionin blue, for 12 hours.

3. Wash rapidly in water.

4. Differentiate carefully in absolute alcohol, or in aniline-alcohol (1-10).

5. Carbol xylol; xylol (quickly); balsam.

Nissl’s granules are blue. This method is easy, and the best for general work.

B. Nissl’s Method.

1. Fix in 96 per cent alcohol for 2-5 days; mount tissue in gum arabic on block; harden in 96 per cent alcohol; cut; place sections in 96 per cent alcohol.

2. Stain in methylene-blue soap mixture (methylene-blue B 3.75 grms., Venetian soap shavings 1.75 grms., water 1,000 cc. Shake well. Keep 3 months before using. Shake and filter before using.), warming, until bubbles arise.

3. Differentiate in aniline alcohol (aniline oil 10 parts, 96 per cent alcohol 90 parts) very rapidly.

4. Arrange section on slide; dry with blotting-paper; cover with cajuput oil.

5. Blot; wash off oil with benzene.

6. Remove benzene; mount the wet section in xylol colophonium, slightly warming; press on cover, and remove excess of colophonium.

Nuclei of ganglion cells light blue: granules dark blue. Toluidin blue, thionin, dahlia, Bismarck brown or neutral red may be used instead of methylene-blue, and often give better results.

2. METHODS FOR STAINING MYELIN SHEATHS.

A. Weigert’s Method.

1. Fix in formol 2-3 days.

2. Primary mordant (potassium bichromate 5 grms., fluorchrom 2.5 grms., water 100 cc.: boil and filter) 4-6 days.

3. Without washing after-harden in graded alcohols, in the dark.

4. Imbed in celloidin.

5. Secondary mordant (neutral copper acetate 5 grms., fluorchrom 2.5 grms., water 100 cc., boil and add 36 per cent acetic 5 cc.) for 1 day at 37°C.

6. Transfer to 70-80 per cent alcohol.

7. Cut.

8. Stain in Weigert’s iron-hæmatoxylin, 24 hours.

9. Wash in water, 30 minutes or longer.

10. Differentiate in borax-potassium ferricyanide (potassium ferricyanide 2.5 grms., borax 2 grms., water 100 cc.) until the gray substance appears yellow to white. Control under microscope.

11. Wash thoroughly in water.

12. Dehydrate in absolute alcohol; clear in xylol; mount in balsam.

Myelinated fibres, blue-black, upon a colorless or light yellow background; red blood cells may be blue-black. Weigert’s method is better than any of the numerous modifications.

B. Orr’s Osmic-Acid Method.

1. Place fresh tissue from cerebral cortex or cord, not more than ⅛ inch in thickness, in 1 per cent acetic, 2 cc., and 2 per cent osmic acid 8 cc., for 48 hours. If mixture is darkened at end of 24 hours, renew.

2. Transfer to 10 per cent formol for 3 days, in order to complete reduction and hardening.

3. Imbed in celloidin or paraffin; cut.

4. Remove paraffin; alcohol; water; differentiate in ⅛-1/12 per cent potassium permanganate.

5. Transfer to a 1 per cent oxalic acid solution, until sections become yellowish-green.

6. Wash; dehydrate; xylol; balsam.

Nerves and fat black: tissue yellowish-green. A very reliable method.

3. STAINING OF AXIS CYLINDERS.

Stain with Van Gieson’s (red), Mallory’s aniline blue method (red), lithium carmine (red), or

Williamson’s Modification of Bielschowsky’s Method.

1. Fix in Müller’s; imbed; cut.

2. Place sections in 10 cc. of tap water containing a few drops of formalin, 5 minutes.

3. Wash in water.

4. Place in the following silver bath 5-10 minutes: 3 drops of liq. ammoniæ (B.P.) are dropped into a test tube. Add 10 per cent silver nitrate solution, drop by drop, until a brownish precipitate is formed. Dissolve the latter by adding ammonia, drop by drop, until the fluid is quite clear. Add tap water up to 10 cc.

5. Wash thoroughly in water.

6. Transfer to the dilute formol solution until sections become grayish-black (1-3 minutes).

7. Place in the following solution for a few minutes: To 10 cc. of water add 2 drops of 1 per cent watery solution of chloride of gold, a few drops of saturated borax solution, and a few drops of a 10 per cent solution of potassium carbonate.

8. Transfer to a 10 per cent aqueous solution of sodium hyposulphite for a few minutes.

9. Wash in water; dehydrate in alcohol; clear in oil of cajuput; xylol; balsam.

Axis-cylinders, intracellular fibrils and Golgi’s network are stained.

4. STAINING OF NEURO-FIBRILLAR STRUCTURES.

These are stained by Bielschowsky’s method, and by acid fuchsin after fixation with osmic acid. The special methods (Apathy’s gold method, Bethe’s molybdic method, the silver methods of Ramen y Cajal and Robertson) have little practical application in pathologic work, and are used chiefly in the study of normal histology.

5. THE STAINING OF NEUROGLIA.

A. Weigert’s Method.

1. Fix small pieces of fresh tissue in 10 per cent formol for 24 hours.

2. Mordant. 8 days at room temperature (4 days at 37°C.) in copper acetate 5 grms., fluorchrom 2.5 grms., acetic acid 5 cc., water 100 cc.

Or, harden and mordant at the same time in 9 parts of the copper mordant, and 1 part of commercial formol for 8 days, changing on the second day, and once again later.

3. Wash in water: after-harden in alcohol; imbed in celloidin; cut.

4. Place sections in ⅓ per cent aqueous solution of potassium permanganate.

5. Wash in two changes of water.

6. Place in the following reducing mixture, 2-4 hours: Chromogen 5 grms., formic acid (sp. gr. 1.2) 5 cc., water 100 cc.; filter; to 90 cc. add 10 cc. of 10 per cent sodium sulphite solution just before using.

7. Wash twice in water.

8. Place sections in 5 per cent carefully filtered aqueous chromogen solution 10-12 hours. (The glia fibres become darker, and a yellowish contrast is obtained for the ganglion and ependymal cells and thicker axis cylinders. Connective-tissue is stained red.)

9. Wash in water.

10. Place section on a slide freshly cleaned with alcohol; dry with filter paper; stain in the following mixture for about 30 seconds: Saturated solution of methyl violet in 70-80 per cent alcohol 100 cc., oxalic acid 5 per cent solution, 5 cc.

11. Remove excess of stain; dry with filter paper; cover slide with saturated solution of iodine in 5 per cent potassium-iodide solution, 30 seconds.

12. Remove iodine solution; dry with filter paper; differentiate in a mixture of equal parts aniline oil and xylol until no more heavy clouds of stain are given off. Control under microscope.

13. Dry section with filter-paper; add xylol; blot; repeat three times.

14. Mount in balsam or turpentine colophonium.

Neuroglia fibres and nuclei, blue; connective-tissue, blue-violet; thicker myelin sheaths, ganglion and ependymal cells, yellowish. This is the best method, none of the modifications giving as good results. No method, however, will stain every neuroglia-fibre.

B. Mallory’s Neuroglia Method.

1. Fix small pieces in 10 per cent formol, 4 days.

2. After-harden in saturated watery picric solution 4-8 days: or combine 1 and 2 by using formol 10 cc. with 90 cc. saturated picric acid solution.

3. Place in a 5 per cent aqueous solution of ammonium bichromate, 4-7 days at 37°C., changing solution 011 second day; or 3-4 weeks at room-temperature.

4. Without washing, harden in alcohol; imbed in celloidin or paraffin; cut.

5. Place sections in ¼ per cent aqueous solution of potassium permanganate, 15-30 minutes.

6. Wash in water.

7. Immerse in 5 per cent aqueous oxalic acid, 5-30 minutes.

8. Wash in several changes of water.

9. Stain in following solution, 1-several days: Hæmatoxylin 0.1 grm., 10 per cent phosphotungstic acid 20 cc., hydrogen peroxide 0.2 cc., water 80 cc.

10. Wash rapidly in water.

11. Differentiate in freshly prepared 30 per cent alcoholic solution of ferric chloride, 5-20 minutes.

12. Wash in water.

13. Dehydrate in 95 per cent and absolute alcohol or blot with xylol.

14. Clear in xylol; balsam.

When Zenker’s fixation is used, omit 2 and 3, and after cutting sections treat with Lugol’s to remove mercury and then with 95 per cent alcohol to wash out iodine; then wash in water and proceed with 5.

Neuroglia, nuclei and fibrin, dark-blue; all else is pale yellow or gray. If the differentiation in 11 is omitted, the axis-cylinders and ganglion-cells are rose-pink; the connective-tissues, dark red-pink.

6. COMBINED STAINING OF SEVERAL NERVOUS STRUCTURES.

Various methods of impregnation with silver, gold or lead are used in histologic work, the Golgi methods and their modifications in particular. They have but little application in pathologic work, and for that reason are omitted here, as is also a consideration of Ehrlich’s vital methylene-blue method and its modifications. Full details of these methods can be found in laboratory textbooks on histology.

7. METHODS FOR THE DEMONSTRATION OF NERVE-DEGENERATION.

A. Marchi’s Method.

1. Harden small fresh pieces of tissue in Müller’s fluid for at least 8 days. Handle tissue very carefully to prevent mechanical injury. The tissue may be placed first in formol, and then later transferred to Müller’s fluid.

2. Place in freshly prepared Marchi’s fluid (Müller’s fluid 2 parts, 1 per cent osmic acid solution 1 part) for about 8 days in the incubator at 37°C. The brain requires a longer time. When the mixture loses the osmic acid smell renew it.

3. Wash in running water, 24 hours.

4. Harden in graded alcohols.

5. Imbed in celloidin; cut; dehydrate; clear; mount.

Degenerated nervous tissue (fat) is black: all else brownish gray. Contrast stain in Van Gieson’s, lithium carmine, etc. This method is good for the demonstration of early degenerations.

B. Donaggio’s Methods for Early Degeneration.

Method I—

1. Fix in Müller’s fluid or in 4 per cent potassium-bichromate solution. The tissue may remain in the fluid for any length of time.

2. Transfer directly, without washing, to alcohol. Dehydrate; imbed in celloidin; cut sections 20-30 microns. Place sections in distilled water for a few seconds.

3. Transfer to the following mixture for 10-20 minutes: To 20 per cent solution of ammoniated chloride of tin add an equal amount of 1 per cent aqueous hæmatoxylin. Allow to stand for five days. Keep in the dark, and in a cool place.

4. Wash rapidly in distilled water.

5. Differentiate in Pal’s solution (oxalic acid 0.5 grm., potassium sulphite 0.5 grm., water 100 cc.) until the normal fibres are entirely decolorized.

6. Dehydrate; xylol; neutral balsam.

Degenerated fibres blue; normal, decolorized.

Method II—

1. Fix in Müller’s fluid: imbed as in Method I.

2. Place sections in 0.5-1 per cent aqueous hæmatoxylin solution, 10-20 minutes.

3. Transfer directly to a saturated aqueous solution of neutral acetate of copper, 30 minutes. Renew copper solution once.

4. Decolorize as in Method I.

5. Wash rapidly in distilled water.

6. Dehydrate in graded alcohols; xylol; balsam.

Degenerated fibres black; normal fibres unstained, except for a narrow circle at periphery.

Method III—

1. Fix and imbed as in Method I.

2. Stain in 0.5-1 per cent aqueous hæmaloxylin solution, 10-20 minutes.

3. Transfer directly to 10-20 per cent solution of perchloride of iron. The section becomes black. After a few seconds they lose their color. If washed in water, they regain their color.

4. Without washing, differentiate in acid alcohol (0.75 cc. HCl in 100 cc. alcohol).

5. Dehydrate in absolute alcohol; xylol; balsam.

Degenerated fibres appear as small black streaks or circular areas.

C. Staining of Fat-granule Cells.

Fix in Flemming’s or Marchi’s mixtures; or in formol, staining with sudan III or scharlach R. The tissues may be examined also in the fresh condition.

D. Old Degenerations.

Use Weigert’s myelin method to show absence of myelinated fibres. Van Gieson’s method stains the neuroglia of degenerated areas a deep red; it is very useful combined with Weigert’s myelin stain. Weigert’s neuroglia stain may be used to demonstrate the relative parts played by neuroglia and connective-tissue in the formation of sclerotic patches. When Weigert’s iron-hæmatoxylin is used with Van Gieson’s the neuroglia remains unstained, while the connective-tissue stains red. With other hæmatoxylins the neuroglia cannot be sharply differentiated from connective-tissue when stained with Van Gieson’s.

8. PERIPHERAL NERVES.

Use any of the above methods for the staining of myelin sheaths, ganglion cells, axis cylinders, etc. Van Gieson’s is good for the demonstration of connective-tissue increase. For the demonstration of peripheral fibrils and nerve-endings consult textbooks on histology for Golgi methods, Ehrlich’s vital methylene-blue method, and the modifications of May, Drasch, and others.

Platner’s Method.

1. Harden in 25 per cent solution of liq. ferri sesquichlor., 1-5 days.

2. Wash in water, until the addition of KCNS to the water yields no reaction.

3. Place in 75 per cent alcohol containing an excess of di-nitroresorcin, 2-30 days, according to the size of the piece of tissue.

4. Dehydrate in absolute alcohol.

5. Imbed; cut; dehydrate; clear; mount.

Axis cylinders, emerald green. A good method for the rapid demonstration of pathological processes connected with the peripheral nerves.

9. CHROMAFFINIC TISSUES.

Wiesel’s Method.

1. Fix 1-4 days in 10 parts of a 5 per cent potassium bichromate solution, 20 parts of 10 per cent formol, 20 parts distilled water.

2. 1-2 days in 5 per cent potassium bichromate.

3. Wash thoroughly in running water, 24 hours; harden in graded alcohols; imbed in paraffin.

4. Stain sections in a 1 per cent aqueous toluidin-blue or water-blue solution.

5. 5 minutes in tap-water.

6. Stain 20 minutes in a 1 per cent watery safranin solution.

7. 96 per cent, and then absolute alcohol, until the blue color appears.

8. Xylol; balsam.

Chromaffinic cells green; other cells light blue, nuclei red.

X. SKIN.

Skin should be fixed in formol-Müller’s or formol, and should not be left too long in alcohol or xylol. Imbed in celloidin preferably. For general use the ordinary stains suffice; for the study of pigment stain with alum or lithium-carmine. Use Weigert’s elastic-tissue stain for the demonstration of elastic fibrils. The intercellular bridges may be demonstrated by Van Gieson’s (remaining unstained), or by various special staining methods.

Herxheimer’s Method for Epithelial Fibrillae.

1. Harden; imbed; cut.

2. Stain in a saturated watery solution of kresyl-echt-violett.

3. Dehydrate in absolute alcohol; clear in clove oil; balsam.

CHAPTER XXIX.
MICROSCOPIC EXAMINATIONS FOR MEDICOLEGAL PURPOSES.

1. BLOOD. Fresh spots should be scraped off and examined in physiologic salt solution. Older spots are scraped off when possible, or if the spots are on clothing or other fabrics, a piece of the stained portion is cut out, and the scraping or piece of material is placed on a slide in a macerating fluid (30 per cent caustic potash; glycerin 3 parts and concentrated sulphuric acid 1 part; 15 per cent tartaric acid; equal parts alcohol and ether; or Pacini’s fluid [mercuric chloride 1 grm., sodium chloride 2.0 grm., glycerin 100 cc., water 300 cc.]). Even in very old clots or stains some red cells may retain their characteristic form. The process of maceration should be observed directly under the microscope, as the macerating fluid gradually changes the cells after they become loosened. Schmorl advises the following method:—

Moisten a small piece of the fabric in water and stain with hæmatoxylin. Differentiate in acid alcohol, wash thoroughly in water, stain with 1/1000 watery eosin solution, wash in water (3-6 hours), place in alcohol, and then again in water. Tease with great care into fine threads on the slide, and examine the isolated fibres in a drop of glycerin. The red corpuscles are easily recognized; and the nuclei of the white cells or of the red cells of birds and amphibia stand out distinctly. Permanent mounts may be made by passing the specimen through alcohol, xylol, and mounting in balsam. A portion of the stained fabric may be imbedded in celloidin, sections cut, and stained with hæmatoxylin and eosin.

The red cells may be measured by the ocular micrometer. Those of man are somewhat larger than those of other mammals, but the difference in size is so slight that an absolute determination of the source of the blood is not possible from the consideration of size alone. Human corpuscles measure 0.0077 mm. in diameter; the nearest in size are the corpuscles of the dog, rabbit, hog, cow, horse, cat and sheep, in the order given, those of the sheep being smaller. The measurements of a large number of corpuscles must be taken if the question of size is to be considered. When red cells cannot be found the stains must be tested for the presence of blood pigment (Teichmann’s hæmin crystals). A small piece of the scraping or stained fabric is placed upon a slide in glacial acetic acid; the fluid is expressed; and a small crystal of sodium chloride or sodium iodide is added to the fluid. Cover with cover-glass and heat gently until bubbles are given off, and continue carefully until the acetic acid has evaporated. The slower the evaporation the larger the crystals will be. Examine with a ⅙ inch or No. 7 objective. The hæmin crystals are brown or claret-colored, of the shape of rhombic plates single or superimposed. They are insoluble in water, alcohol and ether; dissolve slowly in ammonia, dilute sulphuric and nitric acids, and easily in caustic potash. The form of the crystal and the color determines the diagnosis of the presence of blood, but does not distinguish between human and animal blood. Old stains should be treated with acetic acid 12-24 hours before attempting to obtain the crystals. Contamination of the fabric with fat interferes with the reaction; the fat should be removed first with ether. If the suspected stain occurs on iron, steel, sand, clay or coal, the crystals usually cannot be obtained. Decomposition may also interfere with the reaction. In such cases a solution of the stain in a saturated solution of borax should be made and submitted to spectroscopic examination. The presence of blood-pigment on any substance can be demonstrated by means of the spectroscope or microspectroscope. Within recent years the biologic test for blood and albumins has been turned to medicolegal uses, in the differential diagnosis of human and animal blood or albumins. This test rests upon the principle that in the serum of an animal treated with human blood or albumin there are developed antibodies having a strong specific hæmolytic or precipitating action upon human blood or albumin.

2. Semen. Suspected seminal stains may be tested first by the Florence sperm reaction. A portion of the material is mixed on the slide with a drop of an iodine solution (1.65 iodine, 2.54 potassium iodide, 30 water). Cover and examine with low power. The formation of characteristic long rhombic, brown or violet crystals is evidence of the presence of semen. The reaction is hindered by decomposition. The spermatozoa may also be demonstrated in seminal stains by soaking the fabric, carefully flattened out, in a watch-glass containing dilute hydrochloric acid (1 drop to 40 cc. distilled water), for 5 minutes to 5 hours, according to the age of the stain. The fabric is then removed and gently rubbed on a slide; the smear is covered with a No. 1 cover-slip, and examined with the oil immersion or the highest dry objectives. The smear may be stained by adding neutral red solution to the macerating fluid, or by drying the smear in the air, fixing in the flame, and staining with hæmatoxylin and eosin, neutral carmine or fuchsin. The staining and macerating solutions may be combined in one. The piece of fabric containing the suspected seminal stain is soaked in a mixture of methyl green 1 grm., hydrochloric acid 15 drops, distilled water 350 cc., for 1-6 hours. It is then removed with the forceps and smeared on the slide, examined in the moist state, or dried, fixed and mounted. The head is stained an intense green.

3. Decidua and Foetal Tissues. Clots, curettings and discharges may be examined in the fresh state by teasing; or the material may be fixed in formol, formol-Müller’s or mercuric chloride. The characteristic structures of the decidual cells, chorionic villi and other fœtal tissues are easily recognized and the presence of any one of these determines the occurrence of pregnancy, abortion or child-birth.

4. Hair. The most important medicolegal question concerning hair is the differential diagnosis between animal and human. The following points must be considered:—

1. The cells of the outer layer (cuticle) of the hair are much larger in the majority of animals than in man, and are much more distinctly seen. The cuticle is much more dentate or serrate in the hair of animals than in that of man.

2. The cortical layer in human hair is much thicker than in animal hairs, as compared to the thickness of the medulla; in animal hair the medulla is thicker than the cortical layer.

3. The cellular structure of the medullary substance is indistinct in human hair; in animal hair it is very prominent. In human hair the medulla is often absent, especially in certain portions of the hair, while in animals it is rarely absent, and then only in sharply localized portions. The hair should always be examined throughout its entire length.

Hairs of the beard are usually thickest (0.14-0.15 mm. in diameter), then follow in order of size the pubic hairs of the female, the hairs of the eye-lids, male pubis, male head, and female head (0.06 mm. in diameter). There are, however, marked individual differences, and the hair of infants is much finer than that of older children and adults. The hairs of the new-born are pointed, as are also the hairs of adults that are protected from cutting, rubbing, maceration, etc. Cut hairs are at first blunt; later more rounded. Hairs that have been torn out usually show a bulbous end with the remains of the hair-follicle; fallen hairs have a closed, smooth, atrophic root. When the question arises as to the definite individual from whom certain hairs may have come a very careful comparison of the given hairs with those of the individual concerned must be carried out, as to color, size, thickness of different layers, shape and size of the transverse section, etc. Paraffin imbedding should be employed for this purpose.

CHAPTER XXX.
THE STUDY OF MOUNTED PREPARATIONS.

1. Preparation. Analyze the method of preparation of the specimen, noting-methods of fixation, hardening, imbedding, injection, impregnation, staining, etc. Note the reaction of nucleus, protoplasm, red blood-cells and connective-tissue to the stain. Look for special staining reactions (hyalin. amyloid, mucin, etc.) and metachromasia.

2. Histology. Look for histologic landmarks by which the organ, or part of the body, from which the specimen is taken can be recognized. If such landmarks cannot be found, analyze the section, element by element, until its histologic features are fully noted and recognized. Answer the question, “From what part of the body does the tissue come?” Carry the differential diagnosis as far as possible, with regard to sex, age, side of the body or organ (for example, in the heart decide as to auricle or ventricle, left ventricle or right, etc.).

3. Pathology. Study next the deviations from the normal. Answer these questions: “Is the tissue normal?” “If not, in what respects does it differ from the normal?” Study pathologic conditions with the low power first, then use the high power for the finer details. Consider the histologic features of the organ (capsule, stroma, parenchyma, ducts, etc.) with respect to pathologic changes.

4. Diagnosis. After the pathologic changes have been fully studied, the diagnosis should be formulated. If the pathologic changes found can be correlated as factors in some specific conditions they should not be considered separately, but the specific condition itself would constitute the diagnosis. When the pathologic conditions have no definite relation to each other they should be classified separately. If a section of kidney shows cloudy swelling, congestion, œdema, small-celled infiltration, hæmorrhage and casts, these different conditions would all be included in the diagnosis of an acute parenchymatous degenerative nephritis. If the general picture corresponds to that of tuberculosis, syphilis, neoplasm, etc., the various associated changes need not be considered alone, but the broad and comprehensive diagnosis should be expressed in as concise terms as possible.

Written descriptions of microscopic appearances should be so full and clear that it is possible to make a diagnosis from the word-picture. A full and adequate analysis of the preparation set forth in concise and clear language is of more objective value than the bare diagnosis. Reproductions by means of drawings are also of great value in assisting the memory or in communicating the results of the observation to others. A technique sufficient for ordinary purposes may soon be obtained. Color work is usually easiest, since the majority of sections are stained in color. The camera lucida or Edinger’s drawing apparatus may be of great service in the enlargement and placing of the various elements of the section; with these instruments but little practice is necessary to produce rapid and accurate representations. The close inspection of the preparation during the drawing often reveals features that previously had escaped attention. Many workers object to drawings on the ground of subjectivity, but the same criticism applies even more to word-descriptions. Especial attention must, however, be paid to this danger, both in the case of drawings and word-descriptions. In the case of scientific work the written description is best backed up by photographs, rather than by drawings. Microphotography has the very great advantage of being purely objective, although dishonest photographs are possible. There are, however, certain limitations to microphotography. Differential details are lost in sections; all parts of the section cannot be fairly represented; and focal and color limitations are great disadvantages. I believe, however, that for pure scientific work word-descriptions should be accompanied by objective microphotographs instead of drawings. The microphotographic outfit of Zeiss has already been recommended as the best. It is impossible to take up the subject of microphotographic technique in this book. The reader is referred to the chapter on “Microphotography” in Aschoff and Gaylord’s Atlas, and to the works of Cresbie, Bagshaw, Kaiserling, Neuhauss, and others. (See article on “Mikrophotographic” in the Enzyklopädie der mikroscopischen Tecknik.)