Practical pathology

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.

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.

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.)