  For bacteriological purposes, sections of tissue are most conveniently prepared by either the freezing method or the paraffin method. The latter is decidedly preferable, but as it is of greater importance to demonstrate the bacteria, if such are present, than to preserve the tissue elements unaltered, the "frozen" sections are often of value. Whichever method is selected, it is necessary to take small pieces of the tissue for sectioning,—2 to 5 mm. cubes when possible, but in any case not exceeding half a centimetre in thickness. Post-mortem material should be secured as soon after the death of the animal as possible. The tissue is prepared for cutting by— (a) Fixation; that is, by causing the death of the cellular elements in such a manner that they retain their characteristic shape and form. The fixing fluids in general use are: Absolute alcohol; corrosive sublimate, saturated aqueous solution; corrosive sublimate, Lang's solution (vide page 82); formaldehyde, 4 per cent. aqueous solution. (Of these, Lang's corrosive sublimate solution is decidedly the best all-round "fixative.") (b) Hardening; that is, by rendering the tissue of sufficient consistency to admit of thin slices or "sections" being cut from it. This is effected by passing the tissue successively through alcohols of gradually increasing strength: 30 per cent. alcohol, 50 per cent. alcohol, 75 per cent. alcohol, 90 per cent. alcohol, absolute alcohol. In both these processes a large excess of fluid should always be used. FREEZING METHOD.1. Fixation. Place the pieces of tissue in a wide-mouthed glass bottle and fill with absolute alcohol. Allow the tissues to remain therein for twenty-four hours. 2. Hardening. Remove the alcohol (no longer absolute, as it has taken up water from the tissues) from the bottle and replace it with fresh absolute alcohol. Allow the tissues to remain therein for twenty-four hours. Fig. 71.—Washing tissues. Fig. 71.—Washing tissues. Note.—If not needed for cutting immediately, the hardened tissues can be stored in 75 per cent. alcohol. 3. Remove the alcohol from the tissues by soaking in water from one to two hours. Remove the stopper from the bottle; rest a glass funnel in the open mouth 4. Impregnate the tissues with mucilage for twelve to twenty-four hours, according to size. Transfer the pieces of tissue to a bottle containing sterilised gum mixture. Formula.—
5. Place the tissue on the plate of a freezing microtome (Cathcart's is perhaps the best form), cover and surround with fresh gum mixture; freeze with ether, or for preference, carbon dioxide, and cut sections. 6. Float the sections off the knife into a glass dish containing tepid water and allow them to remain therein for about an hour to dissolve out the gum. (If not required at once, store in 90 per cent. alcohol.) 7. Transfer to a glass capsule containing the selected staining fluid, by means of a section lifter. 8. Transfer the sections in turn to a capsule containing absolute alcohol (to dehydrate) and to one containing xylol or oil of cloves (to clear). 9. Mount in xylol balsam. Alternative Rapid Method.— 1. Cut very small blocks of the tissue. 2. Fix in formalin 10 per cent. aqueous solution (fixation fluid No. 7, page 82) for 24 hours. 3. Transfer block to plate of freezing microtome and freeze with carbon dioxide vapour. 4. Float the sections off the knife into a glass dish of tepid water. 5. Stain the sections in glass capsules containing selected stains. 6. Place the stained section in a dish of clean water and introduce a glass slide obliquely beneath the section; with a mounted needle draw the section on to the slide and hold it there; 7. Drain away as much water as possible from the section. Drop absolute alcohol on to the section from a drop bottle, to dehydrate it. 8. Double a piece of blotting paper and gently press it on the section to dry it. 9. Drop on xylol to clear the section. 10. Place a large drop of xylol balsam on the section and carefully lower a cover-glass on to the balsam. PARAFFIN METHOD.1. Fixation. Place the pieces of tissue, resting on cotton-wool, in a wide-mouthed glass bottle. Pour on a sufficient quantity of the corrosive sublimate fixing fluid; allow the tissue to remain therein for twelve to twenty-four hours according to size. 2. Pour off the fixing fluid and wash thoroughly in running water for twenty minutes to half an hour to remove the excess of corrosive sublimate. Fig. 72.—L-shaped brass moulds. Fig. 72.—L-shaped brass moulds. Fig. 73.—Paraffin kettle. Fig. 73.—Paraffin kettle. 3. Hardening. Place the tissues in each of the following strengths of alcohol in turn for from twelve to twenty-four hours: 50 per cent., 75 per cent., 90 per cent., absolute. 4. Dehydration is effected by transferring the tissues to fresh absolute alcohol. 5. Clearing. Half fill a wide-mouthed bottle with 6. Infiltration. Place the cleared tissues in fresh chloroform with several pieces of paraffin wax and stand in a warm place, such as on the top of the warm incubator. The warmth gradually melts the paraffin and the tissues should remain in the mixture about twenty-four hours. 7. Transfer the tissues to a vessel containing pure melted paraffin. Place this vessel in a paraffin water-bath regulated for 2° C. above the melting-point of the paraffin used, and allow the tissues to soak for some four to six hours to ensure complete impregnation. The paraffin used should have a melting-point of not more than 58° C. For all ordinary purposes 54°C. will be found quite high enough. 8. Imbed in fresh paraffin in a metal (or paper) mould. (a) Arrange a pair of L-shaped pieces of metal on a plate of glass to form a rectangular trough (Fig. 72). (b) Pour fresh melted paraffin into the mould from a special vessel (Fig. 73). (c) Lift the piece of tissue from the paraffin bath and arrange it in the mould. (d) Blow gently on the surface of the paraffin in the mould, and as soon as a film of solid paraffin has formed, carefully lift the glass plate on which the mould is set and lower plate and mould together into a basin of cold water. (e) When the block is cold, break off the metal L's; trim off the excess of paraffin from around the tissue When several pieces of tissue have to be imbedded at one time, shapes of stout copper, 10 cm., 5 cm., and 2.5 cm. square respectively, and 0.75 cm. deep (Fig. 74) will be found extremely useful. These placed upon plates of glass replace the pair of L's in the above process. When the paraffin has set firmly the screw a should be loosened to allow the two halves of the flange b to separate slightly—this facilitates removal of the paraffin block. Fig. 74.—Paraffin mould. Fig. 74.—Paraffin mould. 8. Cement the block on the carrier of a "paraffin" microtome (the Minot, the Jung, or the Cambridge Rocker) with a little melted paraffin. Greater security is obtained if the paraffin around the base of the block is melted by means of a hot metal or glass rod. 9. Cut sections—thin, and if possible in ribbands. Mounting Paraffin Sections.— 1. Place a large drop of 30 per cent. alcohol on the centre of a slide (or cover-slip) and float the section on to the surface of the drop, from a section lifter. 2. Hold the slide in the fingers of one hand and warm cautiously over the flame of a Bunsen burner, touching the under surface of the glass from time to time on the back of the other hand. As soon as the slide feels distinctly warm to the skin, the paraffin section will flatten out and all wrinkles disappear. (The slide with the section floating on it may be rested on the top of the paraffin bath for two or three minutes, instead of warming over the flame as here described.) 3. Cautiously tilt up the slide and blot off the excess of spirit with blotting paper, leaving the section attached to the centre of the slide. 4. Place the slide in a wire rack (Fig. 75), section downward, in the "hot" incubator for twelve to twenty-four hours. At the end of this time the section is firmly adherent to the glass, and is treated during the subsequent steps as a "fixed" cover-glass film preparation. Note.—If large, thick sections have to be manipulated, or if time is of importance or acids are used during the staining process, it is often advisable to add a trace of Mayer's albumin to the alcohol before floating out the section. If this substance is employed, a sojourn of twenty minutes to half an hour in the "hot" incubator will be found ample to ensure firm adhesion of the section to the slide. The albuminous fluid is prepared as follows: Fig. 75.—Section rack. Fig. 75.—Section rack. Mayer's Albumin.— Weigh out
and dissolve in
Add
Mix thoroughly by means of an egg whisk. Filter into a clean bottle. As an alternative method paint a thin layer of Schallibaum's solution on the slide with a camel's hair pencil; lay the section carefully on this film and heat gently to fix the section. Schallibaum's solution:
Keep in a dark blue bottle in a cool place. Staining Paraffin Sections.— 1. Warm paraffin section over the Bunsen flame to soften (but not to melt) the paraffin, then dissolve out the wax with xylol poured on from a drop bottle. 2. Remove xylol by flushing the section with alcohol. 3. If the tissue was originally "fixed" in a corrosive sublimate solution, the section must now be treated with Lugol's iodine solution for two minutes and subsequently immersed in 90 per cent. alcohol to remove all traces of yellow staining. 4. Wash in water. 5. Stain deeply, if using a single stain, as the subsequent processes decolourise. 6. Wash in water, decolourise if necessary. 7. Flood with several changes of absolute alcohol to dehydrate the section. 8. Clear in xylol. (Oil of cloves is not usually employed, as it decolourises the section.) 9. Mount in xylol balsam. SPECIAL STAINING METHODS FOR SECTIONS.Double-staining Carmine and Gram-Weigert.— 1. Prepare the section for staining as above, sections 1 to 3. 2. Stain in lithium carmine (Orth's) or picrocarmine for ten to thirty minutes, in a porcelain staining pot (Fig. 76). 3. Wash in picric acid solution until yellow. At this stage cell nuclei are red, protoplasm is yellow, and bacteria are colourless. Picric acid solution is prepared by mixing
4. Wash in water. 5. Wash in alcohol. 6. Stain in aniline gentian violet. 7. Wash in iodine solution till dark brown or black. 8. Wash in water. 9. Dip in absolute alcohol for a second. 10. Decolourise with aniline oil till no more colour is discharged. Fig. 76.—Staining pot. Fig. 76.—Staining pot. 11. Wash with aniline oil, 2 parts, xylol, 1 part. 12. Clear with xylol. 13. Mount in xylol balsam. Alternative Gram-Weigert Method for Sections.— 1. Fix paraffin section on slide and prepare for staining in the usual manner. 2. Stain in alum carmine for about fifteen minutes. 3. Wash thoroughly in water. 4. Filter aniline gentian violet solution on to the section on the slide and allow to stain about twenty-five minutes. 5. Wash thoroughly in water. 6. Treat with Lugol's iodine until section ceases to become any blacker. 7. Wash thoroughly in water. 8. Treat with a mixture of equal parts of aniline oil and xylol until no more colour comes away. 9. Wash thoroughly with xylol. 10. Decolourise and dehydrate rapidly with absolute alcohol until there remains only a very faint bluish tint. 11. Clear with xylol. 12. Mount in xylol balsam. (Then fibrin and hyaline tissue are stained deep blue, whilst bacteria which "stain Gram" appear of a deep blue-violet colour.) Unna-Pappenheim Method.— Stain.— Weigh out and mix
and dissolve in Carbolic acid 0.5 per cent. aqueous solution 78 c.c. Measure out
Method.— 1. Place tissue in the above stain for ten minutes. 2. Differentiate and dehydrate with absolute alcohol. 3. Clear in xylol. 4. Mount in xylol balsam. To Demonstrate Capsules.— 1. MacConkey's Method.—Stain precisely as for cover-slip films (vide page 100). 2. FriedlÄnder's Method.— Stain.—
Method.— 1. Prepare the sections for staining, secundum artem. 2. Stain sections in the warm (e. g., in the hot incubator) for twenty-four hours. 3. Wash with water. 4. Decolourise lightly with acetic acid, 1 per cent. 5. Dehydrate rapidly with absolute alcohol. 6. Clear with xylol. 7. Mount in xylol balsam. To Demonstrate Acid-fast Bacilli.— 1. Prepare the sections for staining in the usual way. 2. Stain with hÆmatin solution ten to twenty seconds, to obtain a pure nuclear stain; then wash in water. 3. Stain with carbolic fuchsin twenty to thirty minutes at 47°C.; then wash in water. 4. Treat with aniline hydrochlorate, 2 per cent. aqueous solution, for two to five seconds. 5. Decolourise in 75 per cent. alcohol till section appears free from stain—fifteen to thirty minutes. 6. Dehydrate with absolute alcohol. 7. Clear very rapidly with xylol. 8. Mount in xylol balsam. To Demonstrate SpirochÆtes in Tissues. Piridin Method (Levaditi).— 1. Cut slices of tissue 1 mm. thick. 2. Fix in 10 per cent. formalin solution for twenty-four hours. 3. Wash in water for one hour. 4. Place in 96 per cent. alcohol for twenty-four hours. 5. Measure into a dark green or amber bottle 100 c.c. silver nitrate solution 1 per cent., and 10 grammes pyridin puriss. Transfer slices of tissue to this. Stopper and keep at room temperature three hours, then in thermostat at 50° C. for four to six hours. 6. Wash quickly in 10 per cent. pyridin solution. 7. Reduce silver by transferring slices of tissue to following solution for forty-eight hours.
8. Wash well in water. Take through alcohols of increasing strength up to absolute, keeping in each strength for twenty-four hours. 9. Clear, embed, cut very thin sections, mount, remove paraffin, again clear and mount in xylol balsam. The spirochÆtes if present are black and show up against the pale yellow color of the background. Weak carbol fuchsin, neutral red or toluidin blue can also be used to stain the background if desired, after the removal of the paraffin in step 9. To Demonstrate Protozoa in Sections (Leishman).— Reagents required:
1. Mount section, remove paraffin and take into distilled water as usual (vide page 121). 2. Drain off the excess of water. 3. Cover the section with diluted Leishman (1 part stain, 2 parts distilled water) and allow to act for five to ten minutes (until tissue appears a deep blue). 4. Decolourise with acetic acid solution until only the nuclei appear blue (examine the section wet, with low power objective). 5. If the eosin colour is too well marked treat with the caustic soda solution until the desired tint is obtained (as seen with the 1/6-inch objective). 6. Wash with distilled water. 7. Rapidly dehydrate with alcohol. 8. Clear with xylol. 9. Mount in xylol balsam. |
|
