Practical pathology

by Aldred Scott Warthin

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.