CORA sage eeay Bet ae Sth eae Boia a eet wae THE | , + =RKosmell P. Flower Library | is THE GIFT OF Xo | ¢72,ROSWELL P. FLOWE Tt FOR THE USE OF THE N. Y. STATE VETERINARY COLLEGE. “iii 3 1924 104 224 971 METHODS OF PATHOLOGICAL HISTOLOGY METHODS OF PATHOLOGICAL HISTOLOGY BY C. von KAHLDEN ASSISTANT PROFESSOR OF PATHOLOGY IN THE UNIVERSITY OF FREIBURG TRANSLATED AND EDITED BY H. MORLEY FLETCHER, M.A., M.D., Canras, M.R.C.P. CASUALTY PHYSICIAN TO ST. BARTHOLOMEW’S HOSPITAL AND ASSISTANT-DEMONSTRATOR OF PHYSIOLOGY IN THE MEDICAL SCHOOL WITH AN INTRODUCTION BY G. SIMS WOODHEAD, M.D. DIRECTOR OF THE LABORATORIES OF THE CONJOINT BOARD OF THE ROYAL COLLEGES OF PHYSICIANS (LOND.) AND SURGEONS (ENG.), ETC. London MACMILLAN AND CO. AND NEW YORK 1894 & All rights reserved No JO2& RB w/o mw PREFACE I trust that the translation of the third edition (1893) of Pro- fessor von Kahlden’s Technik der histologischen Untersuchen Pathologisch - Anatomischen Prdéparate, which has long been regarded as a standard work on pathological methods in Germany, will be as generally useful in the hands of English patho- logists who are not familiar with the original work. I take this opportunity of expressing my indebtedness to Professor von Kahlden for the courtesy and ready help he has afforded me during the progress of this translation, and especially for his consideration in furnishing me with the proofs of the last edition. To my friend Dr. Rolleston for much help and advice in correcting the proofs, to Dr. Sherrington for several valuable suggestions which I have inserted as footnotes, and to Dr. Sims Woodhead, who has kindly written the Introduction, I must express my deep sense of obligation. H. MORLEY FLETCHER. HaARr.ey STREET, January 1894. INTRODUCTION It is only within comparatively recent years that morbid histology has been considered of sufficient importance to require a knowledge of a special technique in its students. Continental workers were for long far ahead, in this respect, of our morbid histologists, and we then, as now, though in a lesser degree, depended upon them in great part for our knowledge of histo- logical methods. All the earlier methods of hardening, em- bedding, staining, and mounting were worked out by the normal histologist, and many of them were not applied to pathological work for the reason that they were thought to be useful for the preparation of fresh tissues only. Closely following the histologist came the zoologists of the Naples school, who, under Dr. Dohrn, have introduced so many of our present methods for preserving and hardening delicate tissues. Since the one great addition made by morbid histologists, the special methods of staining introduced and improved by Koch, Ehrlich, and Weigert, and their pupils, morbid histo- logists, and especially those working in the region of comparative pathology, have come to see that the best of the ordinary zoological, embryological, and histological methods can with great advantage be applied to morbid histology. The result of this has been that not only in the study of the nervous system and of bacteria in disease, but also in general pathology, the methods now at the command of the morbid histologist are as perfect as those used by other workers in the field of histology. Von Kahlden’s excellent work on “ Methods,” first issued as a supplement to Ziegler’s Allgemeine und specielle Pathologische Anatomie, and latterly in separate form, has for some time been well vill HISTOLOGY and most favourably known in the pathological laboratories in this country, and in introducing this English edition to a wider circle of readers, Dr. Morley Fletcher has conferred a real boon on the busy student of morbid histology who wishes to utilise, with as little expenditure of time and trouble as possible, the many useful methods contained in Von Kahlden’s work. The translation, as those who are acquainted with the original work will find, is well done; but more important still, the translator has been able to give, in the form of footnotes, additions and modifications to the body of the work, the results of his own experience. These additions are many of them of great practical value, and greatly enhance the value of the book, which has by means of these additions been brought well up to date, and rendered, in its completed form, the most reliable and comprehensive work of its kind in our language. GERMAN SIMS WOODHEAD. CONTENTS CHAPTER I PAGE THE MICROSCOPE AND APPARATUS REQUISITE FOR HISTOLOGICAL Work 1 CHAPTER II EXAMINATION oF FrisH TISsvES : ‘ b CHAPTER III METHODS OF HARDENING ‘ . 2 » 12 CHAPTER IV DECALCIFICATION OF TISSUES. Z ‘ e AG CHAPTER V METHODS oF EMBEDDING— @, CELLOIDIN ; a 322 b, PARAFFIN . ‘ ‘i 24 ¢. CELLOIDIN AND PARAFFIN METHODS COMBINED 26 d. OTHER METHODS 5 : : : Z e QF CHAPTER VI INJECTION OF TISSUES . ‘ ‘ ‘ . : . 28 x HISTOLOGY CHAPTER VII PAGE PREPARING AND Mountine SEcTIONS : : 31 a. MIcroToMES . : 3 32 b. SeRIAL SecTION CUTTING i 36 CHAPTER VIII Srarine PROCESSES, ETC. ‘ ‘ ‘ . 39 a. GENERAL Mrrnops ‘ ‘ - 40 b. NucLEAR-STAINING. 3 ‘ . 44 c. DirFusE AND DousLE STAINING ‘ . ‘ ‘ “ 52 d. STAINING IN BULK. . a . 55 e. STAINING KARYOKINETIC FIGURES ‘ . 56 CHAPTER IX EXAMINATION OF DEGENERATED TISSUES, ETC. : , 62 a. NECROSIS x a ‘ ‘ : . 62 b. PIGMENTARY ATROPHY 63 c. CLOUDY SWELLING é 63 d. Fatty DEGENERATION x 63 e. Mucoip DEGENERATION : ‘ 64 f. Coutory DEGENERATION . 65 g. LARDACEOUS DEGENERATION . ’ : 3 65 h. HYALINE DEGENERATION x é : 3 68 7. GLYCOGEN i ‘ a ‘ 4 “ . 68 k. CALCAREOUS INFILTRATION . 69 1, DETECTION OF IRON. ‘ ‘i ‘ . 69 DETECTION OF PHOSPHATES ‘ : . (2 CHAPTER X EXAMINATION OF PROLIFERATING AND INFLAMED TISSUES . “ . 73 CHAPTER XI EXAMINATION OF BAcrERIA— a. BACTERIA IN FLUIpDs . é . . 76 b. COVER-GLASS PREPARATIONS , ‘ : » 99 c. STAINING FLAGELLA . i F 81 CONTENTS ad. STAINING BACTERIA IN SECTIONS e. SECTIONS OF GELATINE CULTURES J. Sporn-STAINING ‘ ‘ 5 ‘ i ‘ ‘ g. SUMMARY OF MzTHODS FoR STAINING DIFFERENT PATHOGENIU Micro-ORGANISMS CHAPTER XII MovuLps AND OTHER FUNGI CHAPTER XII ANIMAL PARASITES CHAPTER XIV MicroscoricaAL EXAMINATION oF SPECIAL TISSUES AND ORGANS— a. BLoop. ‘i 6. HEART AND BLOOD-VESSELS ¢. SPLEEN AND LyMPHATICS d. SrERous MEMBRANES . e. SKIN 3 , J. Mucous MEMBRANES . g. ALIMENTARY CANAL . h. LIVER AND PANCREAS 7. URINARY ORGANS : kh. RESPIRATORY ORGANS AND SPUTUM. 7, CENTRAL Nervous SysTEM m. EYE n. EAR ‘ o. OssEous SYsTEM 3 p. Muscies, TENDONS, AND BURSAE gq. MALE AND FEMALE SEXUAL ORGANS CHAPTER XV MIcROscoPICAL EXAMINATION FoR MeEpIco-LEGAL PURPOSES a. DETECTION OF BLoop 6. EXAMINATION OF Hain c. DETECTION oF SEMINAL STAINS d. DETECTION oF DEcIDUAL REMAINS . xi PAGE 83 87 87 89 107 108 111 124 125 126 126 129 129 130 1382 133 135 157 157 157 159 160 161 161 163 164 165 CHAPTER I THE MICROSCOPE AND APPARATUS REQUISITE FOR HISTOLOGICAL WORK For ordinary work a high power magnifying 300 diameters, and one, or ena epee low, powers are ab olutel necessary. Two wmlemer Pieces, gh a and low. willMBel ononcHinnaelt. Pe ~ ERRATUM. Page 143, line 6, for dehydrate read decolourize. WM. An. oil, immersion lens* is essential for the examination of bacteria. A drop of some oil, or of a mixture of different oils, which has nearly the same refractive index-as glass, is placed between the objective and the upper surface of the cover-glass. In this way the air, which has a different index of refraction, is excluded. A soft silk rag, moistened with benzole, is used to remove the oil from the lens and from the cover-glass, pro- vided that the Canada balsam has set and fixed the cover- - glass. Dust should be removed from the eye-pieces and objectives with a soft linen rag or camel’s-hair brush. Any Canada balsam, cedar wood oil, etc., that may have dried on the surface of the 1 Water immersion lenses, which were the first form introduced, are now obsolete. 54 e CHAPTER I THE MICROSCOPE AND APPARATUS REQUISITE FOR HISTOLOGICAL WORK For ordinary work a high power magnifying 300 diameters, and one, or perhaps two, low powers are absolutely necessary. Two eye-pieces, a high and low, will be enough, the latter being most generally employed. The eye-piece magnifies the image formed by the objective, and not the object itself; any defects in this image therefore become more conspicuous with the higher eye- piece, and with it the illumination is not so good. Apochromatic lenses are an exception to this rule, for much stronger eye-pieces can then be used, and greater magnification of the image produced, without any loss of definition. Microscopes are generally supplied with both concave and plane illuminating mirrors. The concave mirror is the one generally used, while the plane is more adapted for working with ‘low powers. An oil immersion lens’ is essential for the examination of bacteria. A drop of some oil, or of a mixture of different oils, which has nearly the same refractive index-as glass, is placed between the objective and the upper surface of the cover-glass. In this way the air, which has a different index of refraction, is excluded. . 2:0 grms. 25 per cent sulphuric acid - 100:0 ccm. 4. Wash in water. 5. Dry, Canada balsam. Sections are treated with alcohol, xylol, and Canada balsam. Gabbet’s method is the best for practical purposes ; it is very simple, as warming is dispensed with, and decolourising and XI EXAMINATION OF BACTERIA 101 contrast-staining are done in one operation. The two solutions are also very easy to prepare. Besides this the method is a very certain one even for sections. IV. Czaplewski’s Method. It is assumed that some of the tubercle bacilli present have become decolourised by the strong mineral acids employed, and so may escape detection. To obviate this, fluorescin is used instead of mineral acids. The method, which applies chiefly to cover- glass preparations, is as follows :— 1. Stain in warm carbol-fuchsine. 2. Pour off the excess of carbol-fuchsine, and place in a strong alcoholic solution of yellow fluorescin contain- ing an excess of methylene blue, The cover-glass is dipped into this solution six to ten times, allowing the fluid to flow off it again slowly each time. 3. Stain in a strong alcoholic solution of methylene blue. 4. Wash rapidly in water, dry; Canada balsam. Ehrlich’s and Nielsen’s methods are especially useful in the case of sections containing quite isolated tubercle bacilli, as the staining may be prolonged if desired. Leprosy Bacillus. This closely resembles the tubercle bacillus, and is stained by the same methods. - It differs from it in the fact that it stains much more quickly, but the colour is apt to fade. With sections Gabbet’s method gives good results, and according to my experience, fairly permanent preparations. Leprosy bacilli may be distinguished from tubercle bacilli by the fact that they stain with simple aqueous solutions. Baumgarten’s Method. (1) Stain for six to seven minutes in a dilute alcoholic solution of fuchsine (five drops of a strong alcoholic solution to a large watch glass of water). (2) Decolourise for a quarter minute in acid-alcohol (nitric acid 1, to alcohol 10). (3) Wash in water. (4) Stain in methylene blue; wash in water, etc.; Canada balsam. 102 HISTOLOGY CHAP, The leprosy bacilli are seen as red rods on a blue ground, © while the tubercle bacilli are not stained in the short time employed in this method. As shown by Lustgarten, leprosy bacilli can be differentiated from tubercle bacilli by the fact that leprosy bacilli stained in aniline-water gentian violet or aniline-water fuchsine do not decolourise so readily as tubercle bacilli in a 1 per cent solution of sodium hypochlorite.’ Cholera Bacillus. The cholera bacillus stains very readily in strong aqueous solutions of fuchsine. Ten minutes should be allowed for the staining. It is decolourised by Gram’s method. Sections may be stained with fuchsine or methylene blue. Spirillum of Relapsing Fever (Obermeier). Giinther has devised a method for staining this spirillum in blood. The hemoglobin is removed from the red corpuscles which do not stain, so that the stained spirilla are better defined. 1. The cover-glass is drawn three times through a flame, or better, is heated for five minutes in an oven at 75° C. 2. Wash for ten seconds in 5 per cent acetic acid. 3. Remove the. acetic acid, first by blowing on the cover- glass through a glass tube, and then by holding the cover-glass, with the film, on the under surface, over strong liquor ammonie for several seconds. 4. Stain in aqueous solutions of the aniline dyes. 5. Wash in water, dry; Canada balsam. Nikiforoff’s Method. (1) Harden for twenty-four hours in the following mixture :— 5 per cent aqueous solution of potassium bi- chromate : ‘ ‘ . { equal Saturated solution of corrosive sublimate in ( parts. 0°6 NaCl solution : : (2) Continue the hardening in warm 70, 85, 95 per cent alcohol. 7 An excellent summary of the various methods for staining B. Lepra and a critique on the differentiation of this from B. Tuberculosis are given by C, Slater, Quart. Journ. Micros. Science, 1891.—Ep. XI EXAMINATION OF BACTERIA 103 (3) Embed in paraffin. | (4) Stain for twenty-four hours in the following mixture:— Alcoholic solution of tropzolin . . 5 com. Strong aqueous solution of methylene blue 10 ccm. Caustic potash solution (1: 1000) . 2 drops. (5) Wash in water. (6) Dip two to three times into a mixture of equal parts of absolute alcohol and ether. (7) Bergamot oil, xylol, Canada balsam. Actinomyces. In examining pus for actinomyces the characteristic whitish granules must be sought for. The pus should be spread out on a glass slide and examined on a dark surface. If granules are found, they should be carefully crushed between a cover-glass and a slide, and on microscopical examination the mycelia and pear- shaped processes or clubs can readily be made out. Cover-glass preparations can be made and stained in the same way as sections. A number of methods have been described for staining actinomyces in sections, but they all have the same objection, that they do not invariably give good results, different fungus masses in the same section often staining quite differently. For simply demonstrating the fungus in sections, double-staining with hematoxylin and carmine or eosine does very well, the masses being sufficiently clearly differentiated by their red colour from the blue tissue. Weigert’s method for staining bacteria (vide p. 85) with previous staining in lithium carmine gives excellent results. The mycelia are better defined by this than by any other method. Sharp differentiation of the mycelia from the clubs can be obtained by the two following combinations of Weigert’s method, and the other parts of the tissue also become well stained. I. Cover-glass Preparations. (1) Smear the contents of the follicle on a cover-glass; dry in the air; draw through a flame. (2) Stain in aniline-water saffranine for twenty-four hours. (3) Wash in water for a short time. 104 HISTOLOGY CHAP. (4) Stain for five minutes in a saturated aniline-water solution of gentian violet. (5) Wash quickly in 0°6 per cent salt solution. (6) Dry between blotting-paper. (7) Transfer for two minutes to iodine solution (iodine 1, KI 2, water 300). (8) Dry between blotting-paper. (9) Decolourise until the preparation ceases to lose colour in aniline oil, which should be repeatedly changed. (10) Remove the aniline oil with xylol; Canada balsam. The mycelia stain blue, the clubs brownish red. In _ this method gentian violet is always used as the second stain. II. Sections. (1) Stain in aniline-water saffranine for twenty-four to forty- eight hours. (2) Wash thoroughly in water. (3) Stain in hematoxylin for a half to one minute. (4) Wash thoroughly in water. (5) Stain for a half to three hours in a saturated solution of gentian violet in aniline water. (6) Wash in 0°6 per cent salt solution. (7) Treat with iodine solution (1:2:300) for two to five _ ainutes. (8) Dry on the lifter with blotting-paper. (9) Decolourise in aniline oil until the section ceases to lose colour. (10) Stain in an aniline oil solution of eosine for a half-hour. (11) Remove the aniline oil with xylol; Canada balsam. The fungus masses are stained dark blue, the clubs brownish red, the nuclei violet, and the protoplasm of the cells rose-red. Gram’s method with twenty-four hours’ staining also gives good results, but has a disadvantage as compared with Weigert’s method, in that the absolute alcohol dissolves the celloidin, and the follicles drop out to a greater or less extent. Bostroem recommends the following instead of Weigert’s or Gram’s method :— 1. Stain in aniline-water gentian violet. 2. Transfer to Weigert’s picro-carmine without previous washing. XI EXAMINATION OF BACTERIA 105 3. Thoroughly wash in water. 4. Transfer to alcohol, and wash out the gentian violet until the sections become reddish yellow. The mycelia of the actinomyces follicles remain colourless, the clubs stain red, and the surrounding tissue reddish yellow. Good sections can be obtained of individual nodules by care- fully embedding in celloidin. Weigert has published a method of staining with orseille. 1. Stain for one hour in a dark-red solution of orseille in the following mixture :— Absolute alcohol. : . 20:0 ecm. Acetic acid , 5:0 Distilled water : ; 40:0 ” . Wash in alcohol. Stain in a 1 per cent aqueous solution of gentian violet. . Wash in alcohol. . Xylol, Canada balsam. or HS 99 bo The nuclei of the cells are stained a bluish violet, connective tissue orange, the inner part of the ray fungus pale blue, and the outer part ruby red. Commercial orseille should be left for some days exposed to the air, so that the ammonia it contains shall pass off. Israel states that the clubs of actinomyces are stained a Burgundy red colour by leaving the sections for some hours in a concentrated solution of orcein in water acidified with acetic acid. Baranski’s Method. 1. Stain the cover-glass preparations or sections for two to three minutes with picro-carmine. 2. Wash for a short time in water. 3. Dry with filter-paper or dehydrate with alcohol. 4, Canada balsam. The tissue stains red, the centre of the fungus yellow. The clubs do not stain deeply. 106 HISTOLOGY CHAP. XI Flormann recommends the following process, which closely resembles Kiihne’s modification of Gram’s method (vide p. 86). 1. Stain for five minutes in this mixture :— A strong alcoholic solution of methylene blue 1 part. Water . . ‘ ; . 2 parts. 1 per cent aqueous solution of ammonium carbonate . : ‘ Se <22F) 5 2. Wash for ten minutes in a large volume of water. 3. Place for five minutes in an iodine solution (iodine 1, KI 2, water 300). 4. Wash thoroughly in water. 5. Decolourise by leaving for twenty minutes in an alcoholic solution of fluorescin (one in fifty), which should be changed once. Wash out the fluorescin in 95 per cent alcohol. Place in aniline oil for a few minutes. Oil of lavender. Xylol, Canada balsam. SO (00. SaT Os The mycelia become a dark blue colour, and are very beauti- fully differentiated. The clubs are partly light blue and partly colourless. CHAPTER XII MOULD AND OTHER FUNGI THE tissue containing the fungus should be teased out in water, or 0°6 per cent salt solution. In this way the various parts of the fungus can generally be observed. If the tissue is not sufficiently transparent, it may be cleared by treating with 1 to 3 per cent caustic potash or soda. Glycerine and alcohol pro- duce considerable shrinkage of the mycelia. C. Fraenkel’s method of teasing in 50 per cent alcohol, to which a few drops of ammonia have been added, is very useful. The material, teased as finely as possible, is afterwards examined in glycerine, and may be preserved permanently by edging round with varnish. Absolute alcohol is generally employed for hardening tissues containing fungi, as it prevents any further growth taking place. Less shrinkage, on the other hand, takes ’ place in Miiller’s fluid. Vesuvine (Bismarck brown) and methy- lene blue are best for staining, though the various species behave differently with the different aniline dyes. Aspergillus stains with fuchsine, methyl-violet, and vesuvine. Skin fungi or dermatophytes can be examined by Balzer’s method. The infected hairs and scurf, etc., are first freed from fat by treatment with a mixture of alcohol and ether. They are then stained for a few seconds in an aqueous or alcoholic solution of eosine or in aniline dyes, and after dehydration in alcohol are mounted in Canada balsam. The preparations should be ex- amined in 33 per cent caustic potash if they are not to be kept permanently. The various forms of fungi occurring commonly in the stomach are best stained with a dilute solution of Bismarck brown, as they very easily overstain with other aniline dyes. CHAPTER XIII ANIMAL PARASITES A ROUGH examination of these parasites requires no special preparation. Many of them, such as acarus scabiei, acarus fol- liculorum, oxyuris vermicularis, tricocephalus dispar, anchylo- stomum duodenale, trichina spiralis, distoma hepaticum and lanceolatum, may be examined in water without further trouble. The parasite which is being examined should be flattened out by pressing on the cover-glass with the handle of a needle. The ova of nematodes, cestodes, and trematodes should also be exa- mined in water. Trichinae in muscle may be examined by teasing. A small fragment of the muscle should be squeezed between two glass slides, and the translucent film which results examined under a low power for the parasite. 7 Pieces should be taken from the diaphragm and the muscles of the jaw, and preferably portions of muscles near tendons. Sections may be cut with a freezing-microtome with or without embedding in celloidin. The sections should not be cut too thin. Fairly thick sections should be stained with an aqueous solution of methyl-green (1:30). In this way the capsule of the parasite is better defined than in thin sections. Encapsuled and calcified trichinae become transparent on treatment with acids. Protozoa should be treated with fixing and staining re- agents, such as osmic acid, chromic acid, iodine, the aniline dyes, ete. Protozoa in the contents of the intestine should be examined microscopically at first without treatment, or mixed with salt solution. Coccidia in hardened tissues stain well with gentian violet and vesuvine. Double-staining in hematoxylin and _ eosine, 1 CHAP. XIII ANIMAL PARASITES 109 using a rather stronger solution of eosine than usual, and allow- ing it to act for several hours, is also an excellent method. Plasmodia of Malarial Fever. Fresh blood is examined by Plehn’s method. A drop of liquid paraffin is placed on a slide, and also on a cover-glass. A drop of blood is caught on the paraffin on the cover-glass, and this is placed on the slide so that the blood lies between a double layer of paraffin. The warm stage is used during the examination. A weak aqueous solution of methylene blue is best for staining blood preparations, a dilute solution of the stain in sterilised ascitic fluid or in blood serum being very suitable. Eosine may be added to this solution for double- staining. For double - staining, the cover -glass preparation, which has been fixed with absolute alcohol, should be treated for five to six minutes with the following solution :— Strong aqueous solution of methylene blue . 60°0 ccm. 1 per cent solution of eosine in 75 per cent alcohol 20:0 _,, Distilled water . ‘ . 20:0 ~=~,, 20 per cent caustic potaeh sofation . 12 drops. Bignami’s Method for fixing the Tissues :— (1) Leave the tissue in the following solution for a half to several hours :— Corrosive sublimate : 1:0 grm. Sodium chloride : O75, Acetic acid : 0°5 to 1:0 ccm. Water 100°0 ”» (2) Transfer to alcohol containing some tincture of iodine. (3) Absolute alcohol. Magenta red (Dr. Griibler, Leipzig) as a saturated aqueous solution, or dissolved in carbolic acid water, is used for staining, and is followed by washing in absolute alcohol. A mixture of magenta red and aurantia as a saturated alco- holic solution may be used for double-staining. 110 HISTOLOGY CHAP. XIII Taenia. The heads of tapeworms should be examined under a low power in water, salt solution, or glycerine. The scolices of echinococeus should be scraped off with a scalpel from the wall of the cyst, and examined in water or dilute glycerine. The sickle-shaped hooklets may often be seen in teased pre- parations of dead, and even of calcified echinococci. By tearing the cyst the scolices of taenia cysticercus are set free, and by squeezing them under a cover-glass the hooklets and suckers may be made out. By compressing a fresh ripe proglottis of a tapeworm between two glass slides, the branched uterus distended with ova is brought into view. Sections through the wall of an echinococcus cyst made with a razor or a pair of scissors, and examined in water, show very distinctly its characteristic laminated structure. Methylated spirit is used for preserving and hardening. The cysts become brittle in Miiller’s fluid. If portions of the tissue become separated in preparing sections, it should be embedded in celloidin. For staining, nuclear dyes are used, either alone or in conjunction with eosine or carmine. Microscopic preparations may be mounted either in glycerine or Canada balsam. Glycerine is preferable for unstained pre- parations. CHAPTER XIV MICROSCOPIC EXAMINATION OF SPECIAL TISSUES AND ORGANS BLOOD Blood may be examined by the following methods :— (1) In the fresh state, with or without the addition of what is known as a preserving fluid. (2) As cover-glass preparations, with or without fixing the blood corpuscles by means of a special fixing solution, various staining methods being employed afterwards. (3) By cutting sections of a fixed and hardened drop of blood and subsequently staining them. A complete examination entails more than one of these methods. I. Examination of Fresh Blood. The best way of doing this is to place a very small drop of blood on the middle of a cover-glass, and to lay this carefully on a cleaned glass slide. Care must be taken to place the smallest possible quantity of blood on the slide, as the finer details of the blood corpuscles can only be made out satisfactorily in a very thin layer. A good method for obtaining a thin layer is as follows :—A cover-glass is fastened down on a slide with wax smeared round three sides of it. A drop of the blood is allowed to flow under it by capillary action, and the cover-glass is then completely edged round with wax. Blood preparations can be prepared in 1 Von Kahlden is indebted for many points in this chapter to H. F. Miler, Die Methoden der Blutuntersuchung, Centralbl. fiir allg. Path. u. path. Anat. Bd, III. 112 HISTOLOGY CHAP. this way at the bed-side, and examined afterwards at leisure. Plehn’s method of mounting the blood between two drops of paraffin (vide p. 109) may also be employed. As the blood corpuscles rapidly undergo change in distilled water as well as in acids, and part with their hemoglobin very readily, a preserv- ing fluid, i.e. one that has no action on the corpuscles, should be employed. The following fluids can be used for hardening and preserving blood :— (1) Salt solution (physiological). This is very useful. The proper degree of strength, varying from 0°6 to 0°75 per cent, must, according to Bizzozero, be ascertained for each species of animal. (2) Blood serum, lymph, or amniotic fluid from the same species of animal as that under examination. Iodine can be added to the fluid, or a staining solution, such as methyl-green dissolved in 0°6 per cent salt solution, may be run in from the edge of the cover-glass. (3) A sherry-coloured solution of iodine in potassium iodide is very useful for showing up the outlines of the red blood corpuscles. (4) A saturated aqueous solution of picric acid has the same * property. (5) 0°5 per cent silver nitrate solution. (6) Pacini’s Solution :— Corrosive sublimate 1:0 grm. Sodium chloride ; 5 2°0 grms. Distilled water . 200-0 ccm. (7) Hayem’s Solution :— Corrosive sublimate ‘ 0°5 grm. Sodium chloride 10, Sodium sulphate : 5°0 grms. Distilled water : . 200°0 com. } In order to examine human leucocytes for their amceboid movements (e.g. in blood or pus) Sherrington says that it is not necessary to use a warm stage, because for an hour or so they are very active in an ordinary covered preparation at the temperature of the room, provided that the film is—(a) sufficiently thin (the cells, if in contact with both under surface of cover-glass and upper surface of slide, are incited by the mechanical stimulation, and exhibit amcboid movements ; the film of blood is thin enough for this if it appears quite transparent, as though “ laked,” though not really so. If the film is not thin the leucocytes remain spheroids, and motionless) ; (8) protected from evaporation. (Private communication.)—Ep. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 113 The blood is mixed with this solution in the proportion of 1:100. After standing for some time the corpuscles sink to the bottom. In twelve to twehty-four hours the fluid is decanted, and the hardened corpuscles thus obtained are washed and filtered. Eosine may be added directly to the hardening solution.’ The application of these solutions is limited, as permanent preparations cannot be obtained with them, nor do they prevent with certainty structural changes in the cell nuclei. These disadvantages may be avoided by using fixing re- agents, which, however, alter the outlines of the cells. The following fixing solutions may be mentioned :— ‘(1) Acetic acid, preferably 1 per cent. (2) Osmic acid, 1 to 2 per cent. (3) Flemming’s solution (wide p. 56), as modified by several authors by substituting a weaker solution of chromic acid. The staining is done either by adding the stain, such as methyl-green, methyl-violet, saffranine, etc., to the fixing fluid, or by treating sections of the blood subsequently with them (vide infra). II. Cover-glass Preparations. Aqueous solutions tend to remove hemoglobin from the corpuscles, a fact which must be borne in mind in staining cover- glass preparations. Hayem recommends a potassium iodide solution of iodine as a stain which does not dissolve out the hemoglobin. By treat- ing a film of blood with an iodine solution of a rather deep brown colour, the parts containing hemoglobin at once become brown or violet. This is the simplest method for detecting the presence of nucleated red blood corpuscles in mammals. By Ehrlich’s method, the hemoglobin may be deprived of its solubility and tendency to swell by heating the cover-glass 1 The best results are obtained by adding 10 ccm. of blood to 1 ccm. of normal saline solution, containing 5 per cent neutral potassium oxalate. In this way Sherrington succeeds in keeping leucocytes in the living condition for as long as three weeks. The fluid should be kept in a very cool room or cellar.—Ep. I 114 HISTOLOGY — CHAP, film, which has already been dried in the air, for one to several hours on a sheet of copper at 120° to 150° C. The same result may be obtained"more simply by Nikiforoff’s method. The cover-glass film, dried in the air, is placed for two hours in a mixture of equal parts of alcohol (absolute) and ether, and is then dried and stained. Hayem recommends with the same object the action of osmic acid vapour on the dried preparation. In preparing blood films, the smallest possible drop of blood is placed on one cover-glass and covered with another. When the drop has spread out they are carefully drawn apart. It is essential that the cover-glasses should be held by forceps and not by the fingers, because, as Ehrlich showed, even the moisture of the fingers in the manipulation is sufficient to produce a con- siderable alteration in the blood corpuscles. In the examination of blood it is of great importance to be acquainted with the different varieties of white corpuscles which occur in it. According to Ehrlich they are as follows :— (1) Small Lymphocytes.—These are only a little smaller than red blood corpuscles, and have a large, deeply-staining nucleus which occupies the greater part of the cell, so that only a small rim of protoplasm is present around the nucleus. (2) Large Lymphocytes. — These are a further stage in the development of the small lymphocytes. They are two to three times larger than red blood corpuscles, and have also a large nucleus, but this differs from that of the small lymphocytes in being surrounded by a distinct broad mass of protoplasm. (3) Mononuclear elements or transitional forms can be distinguished from large lymphocytes by the fact that the nucleus is not uniformly circular, but has a depression in the middle. (4) Polynuclear Leucocytes.—These are smaller than the mononuclear transitional forms, but are larger than red blood corpuscles. They have either a single branching nucleus, or several deeply-staining nuclei. They constitute about 70 per cent of the white corpuscles in normal blood, and possess the power of amceboid movement. (5) Eosinophile Cells.—The nucleus stains less darkly than in the polynuclear leucocytes. xIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 115 The granules present in the protoplasm (vide infra) stain a deep red colour with eosine. Granules in leucocytes and other cells—Ehrlich has shown further that the white blood corpuscles contain granules in the protoplasm around the nucleus, which exhibit distinctly different reactions towards certain groups of aniline colours; and that according to their behaviour with dyes, different varieties of granules can be distinguished, which he calls a, 8, y, 5, e granules. That this depends on true chemical differences follows from the fact that the various granules exhibit other constant features. (1) In their behaviour towards solvents (water, acids, alcohol, glycerine). (2) In size, shape, and refractive index. (8) In their behaviour to raised temperatures. (4) In the distribution of the granules in the body of the cell. A. The most important are the eosinophile cells—that is to say, cells containing granules which stain deeply with acid aniline dyes, especially with eosine. These are known as a granules. Eosinophile cells may be stained as follows :— (1) Heat the cover-glass preparation for several hours at 120° C. (2) Stain for several hours in Khrlich’s acid hematoxylin and eosine solution (vide p. 45). (3) Wash in water, dry ;.Canada balsam. The nuclei of the lymphocytes and of the polynuclear cells stain darkly; the nuclei of the mononuclear cells bluish gray ; the red blood corpuscles copper red, and the eosinophile granules red, Cover-glass preparations treated by the methods described above may also be stained in the following solution :’— Aurantia Induline + 2:0 grms. of each. Eosine Glycerine, 30°0 ccm. ’ For purposes of simple diagnosis cover-glass preparations of the blood, prepared in the ordinary way (vide p. 79) by passing the film three times through a flame, are made. These are stained first in eosine (a 50 per cent alcoholic solution of eosine diluted with an equal volume of water), and afterwards in methylene blue.—Ep. 116 HISTOLOGY CHAP. Wash in water, dry; Canada balsam. The nuclei stain blue; the eosinophile cells red or reddish black, and the red blood corpuscles copper red. For showing the eosinophile cells alone in cases where nuclear-staining is unnecessary, a simple eosine solution is used. A deep red solution of eosine in glycerine is best for this purpose, followed by washing in water. The presence of these eosinophile cells is, according to Ehrlich, of great importance, who states that they are not increased in number in ordinary acute leucocytosis, but, on the other hand, are very considerably increased in leukeemia. They only occur in small numbers in normal blood. Accord- ing to Ehrlich, the fact that the granules become stained by one of the acid dyes is not sufficient to be certain that they are eosinophile granules, and it is necessary to ascertain their colour reactions with the other acid dyes. To make the diagnosis certain the following solutions should be employed :— (a) A deep red solution of eosine in glycerine. (6) A saturated solution of induline in glycerine. (c) A saturated aqueous solution of orange. (@) An eosine-induline-glycerine solution.’ B. Basophile Granules.—These stain with the ordinary basic aniline dyes (the bacteria stains, such as methylene blue, gentian violet, fuchsine, Bismarck brown, etc.). The y granules | and 6 granules belong to this group. The y granules are also known as “mastzellen” granules. They do not occur in normal human blood, but are very numerous in leukemia. The 6 granules, which also take up basic aniline stains, are found in the mononuclear transitional forms in human blood. The basophile y granules are coarse, the § granules finer. ' The following is the stain employed by Demoor and Massart in their recent work on leucocytes :— : Hematoxylin . 0°90 grm. Eosine , 20 yy Alum . é + 20:0 54 Absolute alcohol Glycerine. } aa 90 ccm. Distilled water The results obtained with this solution vary somewhat according to its age. Before mounting in Canada balsam it is as well to examine the depth of staining in each cover-glass, and to increase this, if necessary, by the addition of eosine or hema- toxylin, as the case may be.—Eb. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 117 Ehrlich recommends the following solution for showing y- granules :— Glacial acetic acid . 12:5 com. Absolute alcohol . ‘ F 50:0, Water . : 100°0 2 Dahlia to almost complete saturation. Stain for several hours, wash in water, and remove excess of the stain in alcohol. Cells containing these granules can be stained in sections. The tissue is hardened in alcohol, stained for twelve hours, and decolourised in alcohol. By Westphal’s method the nucleus can be stained at the same time by using the following solution :— Grenacher’s carmine (pure carmine, 2:0 grms., alum, 5:0 grms., distilled water, 200°0 ccm. This is boiled, filtered, and 1:0 cem. carbolic acid added) . 100 cem. Glycerine : F ; 100 _ , A strong solution of dahlia in absolute alcohol 100 Glacial acetic acid : . ; 20 Wash the cover-glass after staining, allow-it to dry in the air, mount in Canada balsam. The “mastzellen” appear as masses of reddish violet granules, in the middle of each of which is a spot corresponding to the cell nucleus. Nuclei of other cells are stained violet or blue. According to Westphal, impure commercial methyl-green stains the y granules bluish violet and the nuclei green. The 6 granules are stained by leaving the cover-glass for ten minutes in a strong aqueous solution of methylene blue, washing in water, drying, and mounting in Canada balsam. C. Neutrophile Granules or ¢« Granules. These stain with neutral aniline dyes'—that is to say, with those produced by mixing a basic with an acid dye. They occur 1 Kanthack and Hardy state that the so-called neutrophile reaction (7.e. that produced by what were believed to be neutral dyes) is really an eosinophile (oxyphile) reaction, the ‘‘neutral’’ dyes being, as they show, really acid dyes. —Ep. 118 HISTOLOGY CHAP. closely packed together in the polynuclear leucocytes. They are also found in various mononuclear leucocytes in the blood of myelogenous leukemia. Method of Staining. (1) Prepare a cover-glass film by Ehrlich’s method (wide ante). (2) Stain for several minutes in the following mixture :— Saturated aqueous solution of orange i . 1205 parts. Saturated aqueous solution of acid-fuchsine contain- ing 20 per cent of alcohol ; » 125 -,, To this is added— Absolute alcohol ; ‘ « 6 Saturated aqueous solution of tiathylgeeen 5 225 x This solution should not be used until it has been kept for some time. The composition of the solution undergoes changes by filtering, and a precipitate is produced, and it is therefore, in staining, better to take up a drop from the middle of the fluid with a pipette and to place it on the cover-glass. (3) Wash in water, dry; Canada balsam. The hemoglobin stains orange yellow, the nuclei greenish, the eosinophile granules dark gray, the neutrophile granules deep violet. Sharp definition is obtained by the following method :— (1) Stain for five minutes in a mixture of Saturated aqueous solution of acid fuchsine .. . 5 vols. To which is added, while continually stirring, Strong aqueous solution of methylene blue : . I vol. Distilled water : . 5 vols. (2) Wash slightly with satan ‘Draw off the water with blotting-paper. Dry in the air; Canada balsam. The eosinophile cells stain red, the « granules violet. Aronson-Philipp’s mixture also gives very good results. Prepare saturated aqueous solutions of Orange G. Acid-rubin extra. Crystals:of methyl-green. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 119 These solutions, having been clarified by settling, are mixed as follows :— Orange G. solution : : 55 ccm. Acid-rubin _,,__ (acid fuchsine) ‘ 50, Distilled water : . j 100 , Alcohol : ; : : 50g, Methyl-green solution . : é 65, Distilled water : 50 =, Alcohol : 3 ‘ : 12 The solution should stand for some weeks. Stain for several hours. Wash in water, dry in the air; Canada balsam. The nuclei stain green, the hemoglobin orange, the e granules reddish violet, a granules yellowish red. A point of special importance is that these neutrophile or e granules belong to the so-called polynuclear leucocytes, and also to the migratory cells occurring in inflammation. They are found in the greater number of the white corpuscles of normal blood. Pus consists for the most part of these leucocytes with neutrophile granules.’ III. Biondi’s Method for examining Blood by Means of Sections. It consists of cutting sections of blood which has been pre- viously hardened. It can be applied to other fluid collections in the tissues. A drop of blood is placed in 5 ccm. of a 2 per cent osmic acid solution, and broken up in this by shaking. After a time the cells sink to the bottom. In twenty-four hours (no longer) 1 The dyes in general use may be roughly classified as follows :— (1) Basie Dyes— Fuchsine (all kinds except acid-fuchsine). Gentian-violet. Methyl-violet. Methylene blue. Bismarck brown (or vesuvine). (2) Acid Dyes— Acid-fuchsine. Eosine. Picric acid. Hematoxylin crystals in aqueous solution.—Ep. 120 HISTOLOGY CHAP. one to two drops of the osmic acid solution are taken out with a pipette and transferred to 5 ccm. agar-agar, liquid at 33° to 37° C. (Biondi’s agar can be obtained from Konig, 29 Doro- theenstrasse, Berlin.) The osmic acid containing the blood is mixed up with the agar by shaking, and is then poured out into a small paper-box, where it rapidly solidifies. The block of agar is next hardened in 85 per cent spirit, which should be changed several times, and is finally cut between pieces of elder-pith. The sections stain excellently, and the agar, even when deeply stained by aniline dyes, readily decolourises in alcohol. Xylol should not be employed for clearing the sections. Ethereal oils, such as creasote, should be used instead. According to Biondi, better sections are obtained by combin- ing the embedding in agar with soaking in paraffin. The block of agar containing the blood is placed for a day in bergamot oil, and is then transferred directly to paraffin at 45° C. for one to two hours. The block is afterwards allowed to solidify under water. The paraffin must be removed before staining. Similar results are obtained by Rindfleisch’s and by H. F. Miiller’s methods. In Rindfleisch’s method, the blood, which may be fixed by any process, is washed, mixed with a very small quantity of glycerine, and allowed to evaporate on a glass slide, as much as the glycerine present will permit, and at the same time sheltered from dust. A very thin layer of celloidin is then poured over it, which, after it solidifies, can be removed as a thin film. The film is then stained, etc. By H. F. Miiller’s method cover-glass films are prepared of blood or other fluids which do not adhere well to the cover- glass. The cover-glass is placed for a moment on a thin celloidin solution, the excess allowed to run off, and then is dried in the air. It is afterwards stained in the ordinary way by Ehrlich’s method. EXAMINATION OF BLOOD PLATELETS. Blood platelets are flat oval discs, of variable dimensions. They may attain one-third the size of a red blood corpuscle. It must be borne in mind in investigating them that they undergo considerable changes both in size and shape from slight disturbing causes, even exposure to air. The process is as follows :—A large drop of 1 per cent osmic xIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 121 acid solution is placed either on one’s own skin or on the shaved skin of a warm-blooded animal, and the skin is pricked through this drop. The blood by this means does not come into contact with the air. It is then mixed up with the osmic acid, and the platelets can be satisfactorily examined. Instead of osmic acid, which was recommended chiefly by Eberth and Schimmelbusch, the following solution may be em- ployed :— Methyl-violet . 0-01 grms. 0°6 per cent salt solution . 50:0 ecm. Another method for demonstrating blood platelets consists in heating very.rapidly a cover-glass film prepared by Ehrlich’s method (vide p. 113). The blood corpuscles in dried preparations stain diffusely in strong aqueous solutions of methyl-violet, aniline-green, and fuchsine. When, as generally occurs, a granular centre can be differentiated from a homogeneous peripheral portion, the central part stains somewhat more deeply. EXAMINATION AND DETECTION OF FIBRIN. Double-staining with hematoxylin and eosine in sections, cut as thinly as possible, is often very successful, as, for example, to show up the network of fibrin in the red hepatisation of pneu- monia, and in diphtheritic mucous membranes. A better method for examining and staining fibrin is that described by Weigert. It differs from his method of staining bacteria (vide p. 85) only as regards the decolourising fluid used. Weigert’s Method for staining Fibrin, (1) Harden in alcohol. (2) Stain for five to fifteen minutes in a strong aniline- water solution of gentian-violet. (3) Wash in 0°6 per cent salt solution. (4) Dry with blotting-paper on a lifter or glass slide. (5) Two to three minutes on a lifter or glass slide in iodine solution (iodine 1, KI 2, water 100). (6) Dry with blotting-paper. (7) Decolourise in aniline oil, 2 parts. xylol 1 part. (8) Remove the aniline-xylol with xylol. (9) Canada balsam. 122 HISTOLOGY CHAP. In this way the fibrin becomes stained a beautiful blue colour, whilst everything else except bacteria is decolourised. Remains of blood corpuscles, caseous masses, and coagulation necroses in particular, remain unstained. A beautiful double stain is obtained by treating the specimen beforehand with lithium carmine (vide p. 47). Fibrin can be stained in fresh preparations of blood by Lowit’s method. As soon as the blood begins to clot under the cover-glass, 0°6 per cent salt solution is drawn through from the edge until the red blood corpuscles are washed out. The leuco- cytes and platelets remain behind for the most part. Alcohol is next passed through, and the various processes in Weigert’s method are then carried on under the cover-glass. ABNORMAL CONSTITUENTS OF BLOOD. Pigment.—This can be satisfactorily examined by mixing up a small drop of blood in a drop of 0°6 per cent salt solution. Cover-glass preparations can also be prepared either unstained or stained with aniline dyes. Bacteria,— Anthrax bacilli and the spirilla of recurrent fever are the forms of bacteria which occur most frequently in human © ~ blood during life. Anthrax bacilli should be examined either in cover-glass preparations stained by Gram’s method, or in drops of fresh blood. The blood of animals infected with anthrax can also be examined very satisfactorily by the hanging-drop method. In searching for other micro-organisms, especially for cocci, great care must be taken to avoid confusing them with basophile y and 6 granules (vide p. 116), which stain like bacteria with basic aniline dyes. 6 granules are so minute that they cannot be mistaken for any known form of micrococci. The granules of “ mastzellen ” may, on the other hand, closely resemble cocci, but are not generally of such uniforin size as the latter. Staining bacteria in dried (cover-glass) preparations of blood is greatly facilitated by treating them for ten seconds with 1 to 5 per cent acetic acid, thoroughly washing, and then staining. In this way the hemoglobin is removed from the red blood corpuscles, and the bacteria are stained almost alone. Spirilla of recurrent fever may be examined in fresh blood during the febrile attack. They can be recognised by their active move- ments. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 123 Dried preparations should be stained in an aqueous or glycerine solution of Bismarck brown or in Léffler’s methylene blue (see also p. 102).. COUNTING BLOOD CORPUSCLES. The blood must be diluted before counting the corpuscles. A pipette is used for this purpose having a sort of bulb in the middle. This bulb contains exactly 100 times as much as the portion of the tube below it. Blood is first drawn up so as to fill the tube as far as the lower limit of the bulb, and then the diluting fluid is drawn up until the bulb is exactly full. In this way the blood is diluted 100 times. Various fluids have been recommended for dilution. The examination is easiest with the following :— Toison’s Dilution Fluid. Methyl-violet 0:025 grms. Neutral glycerine : . 80:0 com. Distilled water : . 80:0 r To this is added a solution of Sodium chloride. ‘ . 1:0 grm. Sodium sulphate . é . 8:0 grms. Distilled water. i . 80°0 com. The solution is then filtered. In five to ten minutes the white corpuscles become stained violet, and are well differentiated from red blood corpuscles, which stain a greenish colour. Thoma-Zeiss’s counting apparatus (K. Zeiss, Jena) is now filled up with this fluid. The apparatus consists of a chamber 0-1 mm. deep, the floor of which is divided up into 400 squares,’ so that the layer of fluid lying on each square contains zg/gg ccm. 1 The lines engraved on the hematocytometer are often insufficiently distinct. The diluted blood approaches more nearly the refractive index of glass than does air, and therefore, though the lines appeared distinct to the maker of the instru- ment, they tend to give great trouble by disappearing when wanted. The indis- tinctness may be obviated by carefully pencilling over the lines by rubbing a broad- pointed BB pencil lightly across them, or by rubbing in a little of some aniline dye with a piece of silk.—Ep. 124 HISTOLOGY CHAP. The corpuscles are counted in as many squares as possible, not only those in the squares, but also those on the dividing lines of partition. The calculation is made by multiplying the cubic contents, 4000, by the degree of dilution, and by the number of corpuscles counted, and then dividing the product by the number of squares counted. The result obtained gives the number of corpuscles in 1 cmm. of blood. Expressed by a formula the calculation is as follows :— 4000xVxZ when X=number of corpuscles in 1 cmm. of undiluted blood ; V=the extent of dilution, generally 100; Z=number of cor- ~ puscles counted lying on the squares; m=number of squares counted. To estimate the white corpuscles alone, the blood should be diluted by Thoma’s method in the proportion of 1 to 10 with water containing 0°3 per cent glacial acetic acid” This dis- solves the red corpuscles, and the process of counting is thereby considerably facilitated. THE HEART AND BLOOD-VESSELS. The condition of heart muscle can be very well made out by means of teased-out preparations of the fresh tissue, care being taken to tease no more than is necessary. 1 Gower’s hematocytometer, which is very commonly employed in this country for the enumeration of blood corpuscles, will be found described in most physio- logical text-books. It can be obtained from Hawksley, Oxford Street, W. The Zeiss-Thoma hematocytometer is at least twice as accurate as Gower’s instrument. A point to remember in using the Zeiss-Thoma instrument is that the cover-glass must lie absolutely flat on the flat glass rim surrounding the cell. If a particle of dust intervenes, the depth of the cell is increased, and the whole estimation falsified. A practical way of ascertaining whether the cover-glass does lie flat is that, if pressed down firmly by the finger, Newton’s coloured rings (interference phenomena) appear, and the cover-glass adheres by atmospheric pressure. —Ep. 2 This rarely gives good results. Sherrington recommends the following method for getting rid of the red corpuscles. The graduated slide with the diluted blood is placed on the brass plate of a freezing microtome. The ether spray is used until the fluid just freezes. This is readily seen as a sort of wave which passes over it, and it is immediately allowed to thaw. All the red corpuscles will be found to have dis- appeared. The red corpuscles may also be got rid of by heating for five minutes at 60° C. In counting leucocytes it is well to add to the fluid used for diluting the blood (preferably 0°3 per cent solution of neutral potassium oxalate in normal saline) enough methylene blue to make the solution a pale blue. This stains the nuclei gf the leucocytes, and renders them more visible. —Ep. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 125 To differentiate between cloudy swelling and fatty degenera- tion, the reagents described in Chapter I. (p. 10) should be used, acetic acid for the former, and 1 per cent osmic acid for the latter (see also p. 63). Pigmentary degeneration of the cardiac muscle can also be well made out in teased preparations. Miiller’s fluid is the best for hardening, but alcohol may also be used. For cutting sections, the tissue should be embedded in celloidin, and the sections afterwards stained with nuclear dyes. The carmine stains, such as lithium carmine and _ borax carmine, should be chosen when there is any question of pig- ment. Picro-carmine defines the transversely striated muscle fibres very distinctly. Vegetations on valves and elsewhere on the endocardium should be examined in the fresh state for bacteria by means of cover-glass preparations. These should be prepared by finely breaking up the masses on a cover-glass, or by grinding them up in sterilised water with a previously-heated glass rod, and then smearing the fluid on a cover-glass. Pieces to be cut should be embedded in celloidin, though paraffin may also be employed. Sections may be stained by simply treating them with gentian violet and washing in alcohol (comp. p. 83); or Gram’s method may be used, which stains most of the bacteria present in endocarditic vegetations, especially streptococcus and staphy- lococcus pyogenes, and Fraenkel-Weichselbaum’s pneumonococcus. To make out the structure of vegetations, sections should be stained with alum carmine, or with hematoxylin and carmine or eosine. Weigert’s stain for fibrin (vide p. 121) is often of use. The larger blood-vessels are best’ hardened in Miiller’s fluid. The methods given below (p. 126) should be employed for show- ing their elastic tissue. SPLEEN AND LYMPHATIC GLANDS. Miiller’s fluid is the most satisfactory hardening reagent, -though corrosive sublimate may be employed. Sections should be stained with nuclear dyes, though Ehrlich’s methods for staining blood (vide p. 113) can be employed. For examination of the spleen pulp in the fresh condition, 126 HISTOLOGY CHAP, Ehrlich recommends plunging a thick trocar, immediately after death, through the skin into the spleen, and smearing the pulp thus obtained on a cover-glass. Serous Membranes. Miiller’s fluid or alcohol is used for hardening, and the ordinary nuclear dyes for staining. Serous transudations and exudations, when rich in cells, may be examined as fresh preparations in salt solution. If the fluid contains few cells, it should be allowed to settle, . especially if tubercle bacilli or other bacteria are suspected. This separation may be brought about by centrifugalisation. The Skin. Skin should be hardened in Miiller’s fluid or in alcohol, and may be embedded in celloidin. Alum carmine or any other nuclear dye is used for staining. Unna’s method for showing the elastic fibres in skin is as follows :— (1) Harden in absolute alcohol, or in Flemming’s solution, followed by hardening in absolute alcohol. (2) Stain with vesuvine (Bismarck brown). (3) Wash. (4) Stain for twenty-four hours in the following mixture :— Fuchsine ‘ - 0°5 grms, 25 per cent nitric acid . 10:0 ccm. Alcohol a ee ; Distilled water . : ; pe Nee (5) Two to three seconds in 25 per cent nitric acid. (6) Decolourise in dilute acetic acid. (7) Dehydrate rapidly in absolute alcohol, cedar-wood oil, Canada balsam. Herxheimer gives the following method for showing the elastic fibres in skin :— (1) Harden in Miiller’s fluid. (2) Stain for three to five minutes in the following solu- tion :— | i XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 127 Hematoxylin . : 1:0 grm. Solution of lithium carbonate saturated in the cold ; 1:0 cem. Absolute alcohol ; ‘ 20°0_,, Water ; , 20°0 ” (3) Decolourise for five to twenty seconds with ferric chloride solution (B.P.). (4) Wash in water. (5) Alcohol, oil, Canada balsam. The elastic fibres stain a bluish black to black, the surround- ing tissue light blue, the nuclei of connective tissue cells and round cells light blue to blue. The sections may afterwards be stained with Bismarck brown. This method cannot be employed for staining in bulk. Absolute alcohol, picric acid, and Flemming’s solution may also be used for hardening. Miiller’s fiuid is preferable, however, because, decolourisation takes place more readily. Manchot’s method for showing the elastic fibres is as follows:— (1) Harden in Miiller’s fiuid or in alcohol. (2) Stain the section for half a minute in a strong aqueous solution of fuchsine. (3) Remove the excess of the stain in water. (4) Decolourise for one to twelve hours in a sugar solution of the consistency and fluidity of glycerine, to every 10 ecm. of which three to four eae of sulphuric acid are added. (5) Mount the sections in non-acidified sugar solution. The celloidin must be removed from the sections before staining. Unna-Taenzer’s method is very useful. (1) The tissue should be hardened in alcohol, although Miiller’s fluid can also be used. (2) Stain for six to twelve hours in the following solu- tion :— 1 The following method of G. E. Goldmann for staining elastic fibres in skin gives very good results (Bettriige z. klin. Chir. p. 7, 1893) :— (1) Cut in paraffin. (2) Fix the section on a slide, remove the paraffin with xylol and place first in absolute alcohol, and then in an alcoholic solution of crystal violet for twenty-four hours. (8) Decolourise either by Weigert’s method for staining fibrin (vide p. 121) or in Stroebe’s solution used for differentiating axis cylinders (vide p. 139). If the former solution be used the aniline-xylol mixture should contain less aniline than that given.—Ep. 128 HISTOLOGY CHAP. Orcein : : ‘ 0°5 grm. Absolute alcohol. . 40°0 cem. Distilled water 3 i 20:0 _ ,, Hydrochloric acid . . 20 drops. (3) Decolourise in the following solution :— Strong hydrochloric acid 0-1 ccm. 95 per cent spirit . 200 Distilled water : : 5:0 The demonstration of bacteria in the superficial layers of the skin requires special methods, as the corneous layer possesses the same affinity for basic aniline dyes as bacteria do, so that both are therefore equally affected by overstaining or by decolourisation. Before staining, the scales of epidermis must be freed from fat. This can be done in several ways :— (1) The scales are washed in ether and alcohol, stained in an alcoholic solution of eosine, and examined in 33 per cent caustic potash solution (or the staining can be omitted). (2) The scurf, or the skin itself, is freed from fat in alcohol and ether, stained in an aniline-water solution of fuchsine, washed in acid-aleohol (vide p. 47), dehydrated in alcohol, and mounted in Canada balsam. Double-staining with gentian violet may be employed in addition. By Bizzozero’s method a cover-glass is gently rubbed on the part of the skin to be examined. It is then passed through a flame three times, and afterwards the fat removed by alcohol and ether. It is then stained with an aniline dye. After removal of fat, scurf can be examined by the following method :— (a) Transfer from alcohol to a drop of 50 per cent acetic acid or 10 per cent caustic potash on a slide. As soon as the scales have swollen up, a cover-glass is placed over them, and they can be examined. It is often better to treat the scales on the cover- glass with acetic acid, to evaporate off the acetic acid, and then to treat as a cover-glass preparation and stain with Loffler’s methylene blue. (6) The scurf may be examined in glycerine tinged with methylene blue. The bacteria stain blue. Boeck’s method for showing mould fungi in epithelial scales is as follows :— (1) Remove the fat with alcohol and ether. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 129 (2) Stain for a half to several minutes in Sahli’s methylene blue solution, which consists of 5 per cent aqueous solution of borax . 16 parts. Strong aqueous solution of methylene blue 20, Water . s : ; . 24 (3) Transfer for a half to one minute to a weak aqueous solution of resorcin (a few particles of resorcin to a large watch-glassful of water). (4) Leave in alcohol for a few minutes to one hour. (5) Decolourise in a weak solution of hydrogen peroxide. (This decolourisation is occasionally unnecessary.) (6) Alcohol, xylol, Canada balsam. Unna (Monatshefte f. praktische Dermatol., xiii. No. 6, u. 7) gives the following method for detecting bacteria :— The scales, moistened with a drop of acetic acid, are rubbed up to a pulp between two glass slides. The slides are then separated, and dried rapidly over a flame. A mixture of alcohol and ether is then poured on the obliquely-held slide in order to remove the grease. Stain with borax-methylene blue; wash with water. To decolourise more completely, one of the reagents described by Unna (vide pp. 126-128) may be used. MUCOUS MEMBRANES. Harden in Miiller’s fluid. For examination of the epithe- lium, the tissue should be placed in the hardening fluid in as fresh a condition as possible, and should be embedded in celloidin before cutting. Embedding in celloidin is also necessary for the examination of deposits occurring on the mucous membranes. The ordinary nuclear stains are used ; hematoxylin and eosine for double-staining. THE ALIMENTARY CANAL. Miiller’s fluid is the hardening agent chiefly employed. Corrosive sublimate is also suitable. Heidenhain recommends picric acid. For a satisfactory examination of the mucous mem- brane of the stomach and intestine, it is very important that the pieces of tissue be placed as soon as possible in the hardening solution. In the case of the stomach this can readily be done in the cadaver, by filling it with Miiller’s fluid through an india- rubber tube immediately after death. K 130 HISTOLOGY CHAP. The tissue should be embedded in celloidin before cutting. The stains used are the ordinary nuclear dyes; hematoxylin and eosine for double-staining. The Biondi-Ehrlich stain recom- mended by Heidenhain (vide p. 51) is often very useful. The contents of the stomach and intestine generally require considerable dilution before they can be examined under the microscope. This is done by taking up the smallest possible quantity of the fluid with a fine platinum loop, and mixing it up on a glass slide with a drop of water or salt solution. The various constituents of faeces can be separated by centrifugalisation. When blood occurs in the contents of the stomach or intes- tine, a few red blood corpuscles can generally be found ‘in the tarry mass, which render the diagnosis certain. Failing this, the hemin tests should be applied (vide p. 162). : No special methods are required for the detection of other cell constituents, food detritus, etc. Starch granules can be found by the iodine test (compare p. 66). Cover-glass pre- parations should be made for the examination of bacteria. Vesuvine is the best stain for the contents of the stomach, as it picks out very distinctly sarcine and the different yeast fungi. The ordinary aniline dyes should be employed for the intestinal contents. The contents of the intestine should be largely diluted with sterilised water when examining for bacteria. It may be pointed out that a number of bacteria staining blue with iodine solution occur in the intestine. Cover-glass preparations may be made directly from the © intestinal contents when searching for cholera bacilli. Schottelius’s method is better. He recommends diluting the intestinal contents with an equal volume of bouillon, and allowing it to stand uncovered. The cholera bacilli grow most abundantly on the surface, so that preparations taken from this contain comma bacilli in larger numbers than would otherwise be the case. THE LIVER AND PANCREAS. These organs should be hardened in Miiller’s fluid. To detect degenerative changes, the liver cells should be examined in fresh scrapings, with the addition of acetic acid or osmic acid (vide p. 60). Flemming’s solution (cf. p. 56), or Marchi’s solution (vide p. 135), should be used as hardening reagents when examining for degenerative changes. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 131 Preparations hardened in Miiller’s fluid are stained with alum carmine or hematoxylin; for double-staining, with hematoxylin and eosine. Tumours should be embedded in celloidin, as the cells are apt to fall out in cutting sections. Bohm’s Method for showing Bile Capillaries. (1) The pieces of liver, not thicker than 1 cem., are hardened for seventy-two hours in a mixture of Three per cent solution of potassium bichromate, 4 vols. 1 per cent solution of osmic acid ; . 1 vol. (2) Transfer for twenty-four to forty-eight hours to a 0°75 per cent solution of silver nitrate. (3) Wash in distilled water. (4) Harden subsequently in alcohol. Cut. The bile capillaries are darkly stained on a yellowish ground. Bohm’s Method for staining the Fibrous Stroma in the Liver :— (1) Harden pieces of liver, 1 ccm. in size, for forty-eight hours in a half per cent chromic acid solution. (2) Transfer for seventy-two hours to a 0°75 per cent silver nitrate solution. (3) Leave for several hours in distilled water. (4) Harden in alcohol. Cut. The fibrous tissue stains black. The fibrous stroma in preparations hardened in alcohol may also be stained by Oppel’s method. Oppel’s Method. (1) Transfer the pieces of liver from alcohol to a 10 per cent aqueous solution of yellow chromate of potassium for twenty-four hours. (2) Transfer to 0°75 per cent silver nitrate solution. The volume of this solution should be twenty to thirty times that of the pieces of tissue. (3) Change the silver nitrate solution in one hour’s time, and again after two to three hours. (4) Wash for twenty-four hours in distilled water. (5) Transfer to alcohol. Cut. The tissues should not be embedded in paraffin, as this renders them brittle. 132 HISTOLOGY CHAP. URINARY ORGANS. Harden in the same way as the liver and pancreas. Corrosive sublimate also in many cases gives good results. The method of boiling (vide p. 16) should be employed for fixing albuminoid fluids in the glomeruli and tubules. Moderate sized pieces of kidney are plunged in boiling water for one to two minutes, and afterwards hardened in alcohol. The same result is produced by hardening in absolute alcohol, and embedding in celloidin. The same methods are employed for investigating degenerative changes, as in the liver and pancreas. Embedding in celloidin is indispensable in minute examination, as otherwise portions of the secreting tubules always drop out, especially in kidneys which have undergone morbid changes. For staining, the nuclear dyes are used. Urine.— After standing, the sediment is taken up with a pipette, is examined on a slide, or cover-glass preparations may be made. The latter method is used more especially for the detec- tion of bacteria. Tubercle bacilli, in cases of tubercular disease of the urinary tract, are generally only found in small numbers in the urine. The urine should be allowed to stand for twenty-four hours, and a considerable number of cover-glass preparations, six to ten, or even more, made from the sediment, and stained by some of the methods already described. The centrifugal machine may also be used with advantage. Gabbet’s (vide p. 100) method is one of the best on account of its simplicity, and the certain results it gives. In doubtful and important cases it is often useful to stain the cover-glasses in an incubator for twenty-four hours with aniline-water fuchsine.! It is often better, when examining urine for cells, etc., to dilute the sediment with water or salt solution. Acetic acid gives the cells a sharper definition, They may also be rendered more distinct 1 Van Ketel’s method for tubercle bacilli in the urine :— (1) Add carbolic acid to the urine in the proportion of 1 to 20 parts urine. (2) Shake vigorously for at least five minutes. (3) Allow to settle in a conical glass. (4) Draw off some of the sediment with a fine pipette. (5) Make cover-glass preparations by heating in the ordinary way. (6) Dip the cover-glass several times in chloroform (or a mixture of equal parts of alcohol and ether), which renders it transparent. (7) Stain with ordinary carbol-fuchsine. This is a very reliable method, and is also applicable for sputum.—Ep. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 133 by allowing a drop of Loffler’s methylene blue to flow in from the edge of the cover-glass. Permanent specimens may be made from cover-glass pre- parations stained with Loffler’s methylene blue or Bismarck brown. The examination of casts is rendered much easier by suspend- ing the sediment in a dilute iodine solution of a pale yellow colour. Blood in urine can generally be detected by direct micro- scopical examination, as at least a few red corpuscles will probably be seen. In doubtful cases the tests for hemin (vide p. 162) should also be applied. The sediment should be examined fresh for crystals. Acid urate of sodium, which forms a brick-dust coloured deposit when present in large quantities, is amorphous. Uric acid is generally found in the form of “ whetstone” crystals, rhombic plates, or long pointed needles. Urate of ammonium is formed by decomposition of the urine, and occurs as “ hedgehog” crystals. Ammonium-magnesium phosphate (triple phosphates) are oblong crystals (“ coffin-lids ”), Calcium oxalate occurs as square envelope-shaped crystals. Calcium carbonate forms spherical and flat bodies. Bilirubin occurs in both the amorphous form and as yellow rhombic plates. Cystin forms regular hexagonal plates. Tyrosin forms sheaf-like bundles of needles, and leucin spherical bodies. THE RESPIRATORY ORGANS AND SPUTUM. Hardening in Miiller’s fluid is best. The tissues should be hardened in alcohol when fibrin or bacteria are to be sought for. A mixture of gum and glycerine (vide p. 15) is useful when rapid hardening is necessary. To fix inflammatory exudations or cedematous tissue the boiling method (p. 16) should be applied, followed by hardening in strong alcohol. Embedding in celloidin should be employed in all cases in which abnormal contents of the pulmonary alveoli have to be dealt with. The ordinary nuclear stains are employed, and, in addition, the special bacterial stains and Weigert’s stain for fibrin can be used. . Sputum can either be examined undiluted, or after the addition 134 HISTOLOGY CHAP. of salt solution, though this is generally not required. In the examination of sputum it is important to bear in mind that it contains cells from the mouth and pharynx, and that débris of food may also be mixed up with it. Besides salivary corpuscles, squamous epithelium cells from the mouth, round cells, and mucous cells, there may also be found in saliva large cells resembling epithelial cells, but of a rounder shape. They have a large round nucleus, and are considered, although not in all cases, to be desquamated (pulmonary) alveolar epithelium cells. They often contain pigment. They occur in ordinary bronchitis (catarrhal cells). Pigment.—Cells containing pigment granules may originate from hemorrhages, especially from chronic pulmonary congestion due to mitral disease. Red blood corpuscles can often be found as well as the pigment. The pigment resulting from hemorrhage has to be distin- guished from that derived from inspired air. Except in the case of siderosis, adventitious pigment gives no iron reaction with potassium ferrocyanide and hydrochloric acid (p. 71), and is generally black, while blood pigment is brownish red. The difference of colour alone, however, is not diagnostic. Fibrinous casts from the bronchi can be recognised by their characteristic shape. So-called Asthma crystals (Charcot) are long, very pointed octahedra. ; Curschmann’s spirals are ribbon-shaped spiral bodies with a clear thread in the centre. Orystals of the fatty acids occur in fetid bronchitis, gangrene, and abscess of the lung, and in cavities. Many varieties of micro-organisms are found in all sputa. Tubercle bacilli and elastic fibres occur most abundantly in small, caseous, plug-like masses in the sputum. These are the portions, therefore, which should be most carefully examined when searching for either of these abnormal constituents (vide examination of sputum, p. 98). The cover-glass method is employed for the examination of bacteria. Elastic fibres are detected by either treating the fresh sputum with 1 per cent caustic potash, or by boiling it with 10 per cent caustic potash. It is then allowed to settle, and the sediment examined in twelve to twenty-four hours. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 135 THE CENTRAL NERVOUS SYSTEM. Miiller’s fluid is employed almost exclusively for preserving and hardening pieces of tissue from the central nervous system. The spinal cord takes three to four months to harden properly, and the brain four months toa year. The time required for hardening can be considerably shortened by placing it in an incubator. The tissue can be kept in Miiller’s fluid for several years, provided that when the hardening is completed, *the solution is diluted to one-half by the addition of water. The pieces may be kept in spirit for some length of time after this. They should be placed in the preserving solution as soon as possible after removal. It is sometimes advisable not to place the tissue at first in Miiller’s fluid of the ordinary strength, but in one containing only 1 per cent of potassium bichromate. Hardening in Flemming’s solution is very useful in the examination of degenerative changes in nervous tissues. The following method is particularly suitable for detecting slight degenerative changes :— Marchi’s Method. (1) The pieces, which should be as small as possible, are kept in Miiller’s fluid for eight days (or longer if pre- ferred). (2) Place them for six to eight days in the following mix- ture :— Miiller’s fluid é . 2 parts. 1 per cent osmic acid . I part. (3) Wash carefully in water. (4) Harden in alcohol of increasing degrees of strength. (5) Embed in celloidin. In removing the parts from the body great care should be taken to avoid dragging on them. Parts which have undergone degeneration appear black; everything else is light gray. The tissue may be afterwards stained with carmine; preferably with lithium carmine (5 per cent); the staining takes several hours. Weigert’s method of 1 Four weeks is generally best.—Ep. 186 HISTOLOGY CHAP. staining may also be applied, if the preparations are put back again into Miiller’s fluid for a sufficiently long time. Teased preparations of portions of the central nervous system freshly taken from the body can only be prepared with difficulty and are unsatisfactory. The teasing is easier after three to eight days in Miiller’s fluid. There are many different staining methods for the central nervous system. They either stain the nuclei alone, or the axis- cylinders, or the medullary sheaths. I. The simple nuclear stains show all that is required in. many cases, and are specially suitable for small inflammatory foci. II. Nissl’s Method for staining Ganglion Cells. Nissl describes two methods, both of which give good results :— Method a. (1) Harden in alcohol of successively increasing strengths.: The pieces of tissue should be about a half cm. thick. (2) Stain the sections in a strong aqueous solution of fuchsine, which should be heated until vapour arises from the surface, (3) Wash in absolute alcohol for one to two minutes. (4) Clear in oil of cloves until no more colour is given out. (5) Canada balsam. Method b. (1) Harden as in (a). (2) Stain in 0°5 per cent methylene blue solution, which should be heated until it bubbles and begins to boil. (3) When the staining solution has cooled down, wash the sections in the following mixture :— Aniline oil é : 20-0 ccm. 90 per cent alcohol 200-0 _,, This should be continued as long as the colour comes out, and until the white and gray matter are clearly differentiated from each other. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 137 (4) Remove the section on a slide, dry with blotting-paper, and cover it with a drop of origanum oil, which should be allowed to flow off again in a short time. (5) Dry with blotting-paper, and afterwards remove with benzine any origanum oil which remains. (6) Cover the section with a solution of resin in benzine, evaporated down to the consistency of thin Canada balsam. (7) Pass the slide through a flame which ignites the benzine resin, (8) Cover with a cover-glass, and warm slightly until the benzine-resin completely fills the space between slide and cover-glass. III. Axis-cylinder Staining by Van Giesson’s Method. The easiest and most certain method for demonstrating these structures is Van Giesson’s (p. 54). The axis-cylinders stain a brownish-red colour, the medullary substance yellow, the nuclei bluish red, the neuroglia red, the nerve-cells with their processes red, and all sclerosed tissue a brilliant red. The method gives better differentiation and more certain results than were formerly obtained by the use of carmine, and on this account is an excellent general stain. Specimens hardened in alcohol can also be stained by this method, though Miiller’s fluid is preferable, as the differentiation is more marked and the yellow staining of the medullary sheath better defined. IV. Axis-cylinder Staining with Neutral Carmine. Neutral carmine (vide p. 52) often gives good results. It is best to leave the sections in a dilute solution for a long time, up to twenty-four hours, and then to wash them thoroughly. Neutral carmine stains the axis-cylinders, nerve -cells, and interstitial tissue. Degenerated areas appear a deep red colour. The time required for staining varies considerably according to the age of the preparation and the method by which it has been hardened. ‘The staining can be hastened and deepened by placing the sections for ten minutes in a palladium chloride solution (0°01 in 50), and transferring directly to the carmine 138 HISTOLOGY CHAP. solution. Warming also aids the staining. In all cases, however, when time allows, it is best to stain slowly in a dilute solution. The nerve-cells are best stained by neutral carmine, by leaving the sections for twenty-four hours in a light rose-coloured solution. V. Borax Carmine followed by treatment with acid-alcohol (vide p. 47) gives good results. The sections should be left in the stain for a quarter to several hours, though no precise time can be stated. The parts stained are the axis-cylinders, interstitial tissue, nuclei of nerve-cells, and other nucleated structures. Lithium carmine gives similar results, and is employed in the same way. . VI. Czokor’s Cochineal Stain for Axis-cylinders. Cochineal : : ‘ 1:0 grm. Alum ; : ; 10 =~, Water : ; . 100°0 com. This is boiled down to half its volume. Stain for twenty-four hours. Wash in water. The nuclei are stained violet, the axis-cylinders a redder colour. VIL Schmaus’s Carmine Stain for Axis-cylinders. (1) Harden in Miiller’s fluid without washing afterwards. (2) Harden in absolute alcohol in the dark. (3) Stain in the following solution :— Sodium carminate . ; 1-0 grm. Uranium nitrate 5 ; 05 , Water : . 100°0 com. This is rubbed up and dissolved in the water, boiled for half an hour, and filtered when cold. (4) Wash in water, etc. Schmaus also recommends an axis-cylinder stain consisting of a blue-black solution (0°25 grms. in 100-0 com. 50 per cent alcohol), to which some picric acid is added. Stain for one hour. Wash in water. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 139 VIII. Stroebe’s Aniline-blue Stain for Axis-cylinders. To stain the axis-cylinders and medullary substance simul- taneously, Ciaglinski recommended, first staining with saffranine, followed by washing in water, staining with aniline-blue, then washing in water. Stroebe has improved this method by washing in alkaline-alcohol instead of water, after staining with aniline-blue. Stroebe’s Method is as follows :— (1) Harden in Miiller’s fiuid. (2) Stain for a half to one hour in a saturated aqueous solution of aniline-blue. (3) Wash the blue-black sections in water. (4) Place them in a dish containing alcohol, to which twenty to thirty drops of caustic-potash alcohol have. been added. (Caustic potash 1 grm. to alcohol 100 ccm.; allow to stand twenty-four hours, then filter.) The sections are left in this until they become a light brownish-red colour and transparent. One or more minutes are generally sufficient. (5) Transfer for five minutes to distilled water. The sections become light blue again. (6) Contrast-stain for a quarter to half an hour in a satur- ated solution of saffranine diluted to one-half with water. (7) Wash and dehydrate in absolute alcohol. Xylol; Canada balsam. ‘IX. Sahli’s Method. (1) Harden carefully in Miiller’s fluid. (2) Stain the section for ten minutes to several hours in the following mixture :— 50 per cent borax solution : : 16:0 ccm. Saturated aqueous solution of methylene blue 24:0, Distilled water ‘ : ’ 40:0 ,, (3) Wash in water or alcohol until the gray matter has a distinct light blue colour against the dark blue of the white matter. 140 HISTOLOGY CHAP. (4) Clear with cedar-wood oil; Canada balsam. The medul- lary sheaths stain deep blue, the nerve-cells a pale _ greenish colour, and the nuclei of the neuroglia cells © blue. This method is specially adapted for showing at the same time any bacteria which may be present. The specimens cannot be kept for an unlimited time. X. Freud’s gold Chloride Method of staining the axis- cylinders alone. (1) Harden in Miiller’s fluid. (2) Stain for two to five hours in a mixture of equal parts of 1 per cent gold chloride solution and 45 per cent alcohol. (3) Wash in water. (4) Leave for two to three minutes in this solution :— Caustic soda . 10 grm, Distilled water . . 6:0 com. (5) Wash in water. (6) Five to ten minutes in a 10 per cent potassium iodide solution. (7) Wash; alcohol; Canada balsam. Freud’s gold stain does not always prove successful. The axis-cylinders stain dark blue to dark red. The medul- lary layer often stains to a certain extent at the same time. Glass needles should be used for manipulating the sections. There are many modifications of the gold chloride method. Cohnheim, who was the first to employ it for showing the nerves in the corneous layer of the skin, places the sections in 0°5 per cent gold chloride solution, and then for some days in water acidified with acetic acid. The modifications of the method lately recommended consist of the following :— (a) The use of much more dilute solutions in which the sections are left for a correspondingly longer time. (b) The use of the double chloride of gold and potassium or sodium instead of gold chloride. (c) The changing of the gold chloride solution. (d) By reducing with hydrochloric acid, formic acid, or tartaric acid instead of acetic acid. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 141 The gold chloride method is much more successful with fresh than with hardened tissues. XI. Upson’s gold Chloride Method for axis-cylinders. (1) Harden for four to six months in a solution of potassium bichromate in distilled water, which should be repeat- edly changed. The strength of the solution should at first be 1 per cent, afterwards 2 per cent. The hardening process should be carried on in the dark. (2) Wash the hardened pieces of tissue as rapidly as possible in distilled water. Leave them for two to three days in 50 per cent alcohol, prepared by diluting absolute alcohol. This must be repeatedly changed. (3) Leave them in 95 per cent alcohol (prepared from absolute) until they exhibit a distinct green colour ; two to four weeks are generally required for this. (4) Embed in celloidin ; cut. (5) Stain the sections for two hours in the following solu- tion :— Gold chloride ; . 1:0 Hydrochloric acid. : ‘ 2°0 Distilled water : : : 100°0 (6) Wash the sections in distilled water. (7) Transfer to a 10 per cent caustic potash solution for half a minute. (8) Wash in water. (9) Reduce in the following solution, which must be renewed for each section :— Sulphuric acid : : 5 ecm. 3 per cent tincture of jedine 10 to 15 drops. Mix and add one drop of liquor ferri chloridi. (10) Wash in water as soon as the section has turned a rose- red colour. (11) Alcohol; oil of cloves; Canada balsam. The sections should be kept in the dark. All manipulations should be done with platinum or glass needles. Contact with metals must be avoided. 142 HISTOLOGY CHAP, XII. Method of staining Axis-cylinders with Nigrosine. (1) Stain the sections in a strong aqueous solution of nigro- sine for five to ten minutes. (2) Decolourise, first in dilute and then in absolute alcohol. (8) Origanum oil, Canada balsam. This method is useful, owing to its simplicity. It gives a very good general idea of the position of any tracts of degenera- tion that may be present. The following is the best method for staining the medullary sheath of nerve-fibres :— XIII. Weigert’s Hematoxylin Method of staining the Medullary Sheaths.' (1) Harden in Miiller’s fluid. (2) Harden afterwards in alcohol without previously washing in water. (3) Embed in celloidin. (4) Leave the block of celloidin containing the tissue for twenty-four to forty-eight hours in a half-saturated solution of acetate of copper. (5) Twenty-four hours in 70 per cent alcohol. (6) Cut. (7) Stain for fifteen or twenty minutes to twenty-four hours in the following solution :— ? Application of Weigert’s method to sections prepared by the freezing methods. ° W.B. It is difficult in some cases to cut sections of large area, e.g. pons or medulla, by the paraffin or celloidin method. The tissue is hardened in Miiller’s fluid, and afterwards in alcohol, as in (1) and (8) Transfer direct from the alcohol to a saturated aqueous solution of acetate of copper ; leave it in this overnight in an incubator at 40° C. (4) Soak for three to twelve hours, according to the size of the piece, in the fol- lowing solution of mucilage. The mucilage is of B.P. strength, but in place of water a saturated aqueous solution of copper acetate is used. (5) Freeze and cut. (6) Remove the sections from the razor to saturated copper acetate solution, to get rid of the gum, and leave them in this for ten minutes. (7) Transfer to 70 per cent methylated spirit. If allowed to remain in this for twenty-four hours at least, on transference to the Weigert hematoxylin solution the sections turn an ink black in a few seconds. The rest of the process is as usual.—Ep. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 1438 Hematoxylin , js . 1:0 grm. Saturated solution of lithium carbonate 1:0 ecm. Absolute alcohol ag . 100 , Distilled water : i . 60:0 » (8) Wash in a large volume of water. (9) Partially dehydrate in this solution :— Sodium biborate . . . 4:0 grms. Potassium ferricyanide ‘ 50, Distilled water : at - 200°0 ccm. The time required for decolourising cannot be given exactly. It should be continued until the gray matter acquires a distinctly yellow colour. (10) Wash in water. (11) Dehydrate in absolute alcohol. (12) Clear in xylol. (13) Mount in Canada balsam. The pieces of tissue hardened in Miiller’s fluid should still be a brown colour, and should not have become green in the alcohol. If this occurs they should be transferred for a few minutes or-longer to a 4 per cent solution of chromic acid, washed very slightly, and placed in the stain at once. The pieces of tissue or the sections may also be replaced for some time in Miiller’s fluid. Weigert’s method gives very good results; the white matter stains a deep blue-black colour, degenerated parts stain lightly in proportion to the number of nerve-fibres which have under- gone change. Any medullary sheaths remaining among the destroyed nerve-fibres also often take up the stain.’ The time required for staining with hematoxylin varies con- siderably. Spinal cord sections often stain in fifteen to thirty minutes. Sections of the cerebral cortex require up to twenty- four hours, and it is advisable to place the staining fluid for at least part of the time in an incubator. If the sections are taken out of the stain too soon, some of the fibres remain unstained, and deceptive appearances may thus result. It is often advisable to decolourise more slowly by diluting the ferricyanide solution to a half, or even a quarter the strength. 1 Weigert’s method XIII. often shows up very clearly fatty degeneration in cardiac muscle.—-Ep. 144 HISTOLOGY CHAP. If origanum oil is used for clearing, the sections should not be left in it longer than is absolutely necessary, or decolourisation will result. Should it be found desirable to stain a portion of the tissue by some other method, a piece should be cut from the celloidin block before placing it in the copper solution. Instead of placing the entire piece of tissue in the copper solution (4), the separate sections may be treated with the solution after cutting. XIV. Weigert’s Method of staining white matter without differentiation. This modification described by Weigert is as follows :— (1) Harden as in method XIII. (2) Transfer to a mixture of Saturated solution of copper acetate eae ae 10 per cent solution of potassium sodium fertnite ene) and leave it in an incubator for twenty-four hours. (3) Transfer to a mixture of Saturated solution of copper acetate ee Water . ‘ ‘ ean BET, for twenty-four hours in an incubator. (4) Stain in the following solution :— Hematoxylin - 1:0 grm. Absolute alcohol ; é . 10°0 ccm. Saturated solution of lithium carbonate. 1:0 a Distilled water ; : . 90°0 ” (This should be prepared by adding the last two to the alcohol and hematoxylin solution.) (5) Wash in water. (6) Clear in the following mixture :— Aniline oil . ‘ . 2 parts. Xylol , : ; . I part. The sections may be transferred to this mixture straight from 90 per cent alcohol. (7) Mount in xylol-Canada balsam. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 145 If the celloidin is stained too deeply, the sections should be passed through 1 to 5 per cent acetic acid after staining. If the sections themselves are overstained, the potassium ferricyanide decolourising fluid described above should be employed. XV. Pal’s Method. This is a useful modification of Weigert’s method. (1) Harden in Miiller’s fluid, etc. (2) Stain in Weigert’s hematoxylin solution for twenty-four to forty-eight hours, for the last hour in an incubator. (3) Wash in water, to which 1 to 2 per cent of lithium car- bonate has been added. The sections should be stained a deep blue. (4) Transfer the sections for twenty to thirty seconds to a 0°25 per cent solution of permanganate of potash until the gray matter appears yellow. (5) Transfer for a few seconds to the following solution :— ‘Pure oxalic acid ; . 1:0 Potassium sulphite . ‘ . 1:0 Distilled water : : . 200°0 (6) Wash thoroughly in water. (7) Alcohol, xylol, Canada balsam. The advantage of this method is that the separate processes can be carried out much more rapidly. It gives very sharp defi- nition. The tissue between the nerve-fibres being completely decolourised, it may be counter-stained, for which purpose picro- carmine or borax carmine is best adapted. XVI Vassale’s modification of Weigert’s method. (1) Harden in Miiller’s fluid, afterwards in alcohol. (2) Stain for three to five minutes in this solution :— Hematoxylin. ; ; 1:0 grm. Water . : . 100°0 ccm, (Dissolve with the aid of heat.) L 146 HISTOLOGY CHAP. (3) Transfer for three to five minutes to a saturated filtered solution of copper acetate. (4) Wash quickly in water. (5) Transfer the sections to the following solution :— Borax . : ‘ : 2°0 grms. Potassium ferricyanide . ; 25, Water . ; : . 300-0 ccm. (The sections should be kept moving to and fro in this solution.) (6) Wash thoroughly in water. (7) Alcohol, carbolic-acid xylol (1:3), Canada balsam. In order to obtain good results the hematoxylin solution | must be fresh. XVII. Kulschitzky’s Method for staining the medullary sheaths. (1) Harden in Miiller’s fluid, or, better, in Erlicki’s fluid (vide p. 14). (In the latter the tissue should remain for one to two months, and be afterwards hardened in alcohol.) (2) Stain for eighteen to twenty-four hours in the following solution, which should be slightly acidified with acetic acid before using. Hematoxylin ‘ ; . 10 grm. Saturated solution of borax . . 20°0 ccm. Distilled water ; ‘ . 80:0 ,, (3) Wash in water. i Kulschitzky’s staining fluid is yellow at first, but in two to three weeks it turns a deep red, and is then ready for use. Before using it, a few drops of acetic acid should be added to a large watch-glassful of the fluid. The medullated nerve - fibres become stained a deep blue colour ; everything else remains unstained or takes up a faint. yellowish tint. This stain is much improved by leaving the sections for twenty-four hours in a saturated solution of sodium or lithium carbonate. . A more recent method published by Kulschitzky is as fol- lows :— XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 147 (1) Stain for one to three (rarely twenty-four) hours in Hematoxylin (dissolved in absolute alcohol) K : . 1:0 2 per cent acetic aeid : . 100-0 (2) Decolourise in the following solution :— Saturated solution of lithium carbonate 100 ccm. 1 per cent potassium ferricyanide solu- tion ‘ : : x LOo (This generally takes two to three hours.) (3) Wash thoroughly in water. A similar result is obtained by boiling for one to four hours 2 grms. (or more) of carmine in 100 ccm. of a 10 per cent acetic acid solution. The sections are left in this for two to four hours, and are then placed direct in the decolourising fluid given above (2). XVIII. Exner’s Method for showing medullated nerve-fibres. The tissue should be taken from the body as soon as possible, though the method succeeds even when twelve hours have elapsed after death. Pieces of the brain or spinal cord, not more than one-half cm. thick, are placed at once in a 1 per cent osmic acid solution, the volume of which should be at least ten times that of the tissue. ‘ The osmic acid solution is renewed in two days’ time. In five or six days the pieces are washed in water, and are either cut directly after fixing on a cork, or after previous embedding in 1 Kulschitzky gives the following method for staining neuroglia (Anat. Anzeiger Centralblit., 1893 [357]). (1) Harden in the dark for two to three months in Erlicki’s fluid (vide p. 14). (2) Harden subsequently in the dark in absolute alcohol. (3) Embed in paraffin ; cut. 7 (4) Stain in the following solution :— Patent acid rubin 0°25 parts. 2 per cent acetic acid solution 100, 100 ,, Saturated aqueous solution of picric acid Take three to five ccm. of this mixture and add 100 cem. of 96 per cent alcohol. Leave the sections in the stain for half an hour or longer. (5) Wash in two portions of 96 per cent alcohol. The sections should be left in the second for some hours. (6) Transfer to absolute alcohol ; clear ; mount.—Ep. 148 HISTOLOGY CHAP. celloidin. Each section is at once mounted in glycerine, and one drop of ammonia solution (liquor ammonize 1 to water 50) is added to the glycerine on the slide. The medullated nerve-fibres appear a gray or black colour by Exner’s method ; the specimens are, however, not permanent. XIX. Adamkiewicz’s Method. Adamkiewicz has pointed out that saffranine stains the medul- lary sheath of nerve-fibres red, and, on the other hand, the nuclei of the nerve-cells and neuroglia cells and those in the vessel walls violet. The medullary sheaths of degenerated nerve-fibres, however, do not stain. This holds even in the earliest stages of degenera- tion. The method is as follows :— (1) Harden in the various solutions containing chromium salts. (2) Transfer the sections for a short time to water slightly acidified by the addition of a few drops of nitric acid. (3) Stain in a deep red aqueous solution of saffranine No. 0. The sections do not readily overstain in this. (4) Wash in ordinary alcohol. (5) Transfer to absolute alcohol slightly acidified by the addition of nitric acid. (6) Clear in oil of cloves until the colouring matter ceases to come out. (7) Mount in Canada balsam. By this method the medullary sheaths stain yellowish red or red, and the nuclei of the connective tissue cells bluish violet. The results obtained by this method are by no means so con- stant as stated by Adamkiewicz. Nikiforoff’s modification of the method is an improvement. (1) Harden in solutions of chromium salts. (2) Complete the hardening process in alcohol, without previous washing in water. (3) Place the sections, after cutting, directly in alcohol. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 149 (4) Stain for twenty-four hours in a strong aqueous solution of saffranine, or in aniline-water saffranine, or carbolic acid water (5 per cent) saffranine. (5) Wash carefully in alcohol, moving the section to and fro until the gray matter is differentiated from the white by its lighter staining. (6) Transfer to gold chloride solution (1: 500) until the gray matter has a slight violet tinge. (7) Wash carefully in water. (8) Transfer to absolute alcohol until the gray matter is clearly marked out by its violet colour from the white matter which is stained red, (9) Oil of cloves for a short time. (10) Xylol. (11) Canada balsam. XX. Golgi’s Nitrate of Silver Method for showing nerve- cells and their processes. The pieces of tissue, which should be small, are hardened in Miiller’s fluid, and then transferred directly from this to a 0°75 per cent silver nitrate solution, which should be changed in an hour’s time. The pieces may remain as long as desired in the fresh solution. In five to six days the tissue, after being dried slightly, is placed first in dilute, and then in absolute alcohol, and afterwards is cut. Alcohol, oil, Canada balsam. The ganglion cells and their processes stain black, and also the connective-tissue cells. The method is very unreliable, as the nerve-cells rarely all take up the stain. Golgi’s corrosive sub- limate method is an improvement. XXI. Golgi’s Corrosive Sublimate Method for nerve-cells and their processes. The piece of tissue, not more than 0°5 cm. thick, is hardened in Miiller’s fluid, and then placed in 0°25 per cent corrosive sub- limate solution, which should be changed as long as it becomes tinged yellow. Small pieces may be cut in eight to ten days’ time, though it is better to leave them longer in the solution. The sections should be very thoroughly washed. Alcohol, oil, Canada balsam. 150 HISTOLOGY CHAP, By this method also the nerve-cells with their processes stain black, but this method, like the foregoing, is an uncertain one. An improvement suggested by Pal consists in treating the sections afterwards for some minutes with a solution of sodium sulphide (Na,S). Specimens prepared by the corrosive sublimate method are very delicate, and cannot be preserved under a cover-glass. This is also the case with the silver preparations. Sections can be permanently mounted and double-stained by the following method :— i XXII. Obregia’s Method. (1) Corresive sublimate or silver nitrate preparations pre- pared by Golgi’s method are treated with alcohol (of strength not less than 95 per cent). They are then placed for half an hour in the following solution :— 1 per cent ferric chloride solution 8 to 10 drops. Absolute alcohol : . : 10 ccm. The solution should be freshly prepared and exposed to diffused light for half an hour. (Metal needles, etc., should not be used.) The sections should be left in this solution for fifteen to thirty minutes in the dark. (2) Wash quickly in 25 per cent alcohol, then in distilled water. (3) Place in 10 per cent sodium sulphide solution for five to ten minutes (not longer). (4) Wash repeatedly in distilled water. (5) Stain with carmine or hematoxylin by Weigert’s, Pal’s, etc., methods. Mount in Canada balsam, cover with a cover-glass. The following is a slight variation of Kélliker’s modification of Golgi’s method :— XXIII. Ramon y Cajal’s Method. (1) The pieces of fresh tissue are left for twenty-four, thirty- six, or forty-eight hours in the dark in the following — solution :— XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 151 Potassium bichromate : ‘ 3 parts. 1 per cent osmic acid solution 2 DDI gs Distilled water. : . 100 The fluid should be changed after some hours, but may be used again for the preliminary treatment of other pieces of the tissue. (2) Treat with a 0°25 per cent silver nitrate solution for a quarter to half an hour. (3) Transfer to a 0°75 per cent silver nitrate solution for thirty-six to forty-eight hours. (4) Wash in 40 per cent alcohol. (5) Dehydrate quickly in absolute alcohol. (6) Cut by hand between elder-pith, etc. (vide p. 277). Embedding in paraffin and celloidin cannot be employed. Cover-glasses should not be placed on the specimens. These methods, though useful, cannot always be relied upon and the staining is not uniform. Nerve-cells and processes, and all non-medullated nerve-fibres stain deep black; the neuroglia cells and their processes stain reddish black. XXIV. Flechsig’s Modification of Golgi’s Corrosive Sublimate Method. Flechsig devised this method for demonstrating the connection between the processes of the nerve-cells with the reticulum in the gray matter, and further, the connection of the reticulum with medullated nerve-fibres. The first brain examined by this method was hardened in Miiller’s fluid, and was then left for a whole year in 1 per cent corrosive sublimate. (1) Harden in a 2 per cent solution of potassium chromate. (2) Soak in corrosive sublimate solution. (3) Cut. The sections should be placed in 96 per cent alcohol. (4) Stain for three to eight days at 35° C. in the following solution :— 152 HISTOLOGY CHAP, Pure extract of Japanese red wood ; 1 grm. Saturated solution of sodium sulphate : 5 com. 5 o tartaric acid : Bx 3 Absolute alcohol ; , : 10 , Distilled water . ; ; . 900. ,, (5) Transfer each section separately to 3 com. of 4 to 4 per cent solution of potassium permanganate until the fluid loses its colour. (6) Decolourise in the following solution :— Oxalic acid . : 1:0 grm. Potassium sulphite 10 =~, Distilled water . 200°0 ccm. If decolourisation is not complete, repeat again (5) and (6) until the yellow tint has vanished from the section. (7) Transfer the section to a mixture of 1 per cent gold chloride solution, 5 drops. Absolute alcohol . . 20 com. until the mercurial precipitate which was a whitish colour by reflected light becomes a deep black, and the red bundles of nerve-fibres acquire a bluish tint. (8) Wash for a short time in 5 per cent potassium cyanide solution, 1 drop. Distilled water i , 20 ccom. The sections should be floated on the fluid. (9) Dehydrate in absolute alcohol. (10) Clear in pure oil of lavender. (11) Canada balsam. The bundles of nerve-fibres are stained a carmine red; the nerve-cells and processes a deep black colour. XXV. Wolter’s Method for staining axis-cylinders. (1) Harden in the dark for twenty-four hours in the follow- ing mixture :— ' Sulphate of copper Potassium bichromate 50 per cent alcohol to each 100 ccm., to which six drops of acetic acid should be added. } any quantity desired. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 153 (2) Transfer to absolute alcohol for twenty - four hours. Embed in celloidin. Cut. (3) Transfer the sections for twenty-four hours to a mordant composed of Vanadium chloride. : 2 parts. 8 per cent ammonium acetate solution . : F 8 ” (4) Wash for ten minutes in water. (5) Stain for twenty-four hours in an oven in hema- toxyln . , 2°0 grms. Absolute alcohol (sifticiont to dissolve). 3 per cent acetic acid. . 100°0 ccm. (6) Wash in 80 per cent alcohol acidified with hydrochloric acid until the sections acquire a light bluish-red colour. (7) Wash out the acid with dilute alcohol; dehydrate in absolute alcohol; origanum oil; Canada balsam. The nerve-cells and neuroglia stain as well as the nerve- fibre. XXVI. Malbory’s Method for Staining Axis-cylinders. Stain the sections for ten minutes to one hour in the follow- ing solution :— Phospho-molybdic acid é 1 part. Hematoxylin ; ‘ We ays Chloral-hydrate 6 to 8 parts. Water : F . 100 , This solution should be exposed to sunlight for a week, and filtered before using. (1) Wash in 50 per cent alcohol, which should be changed two to three times. (2) Dehydrate, etc. The stained sections should not be left too long in alcohol. The amount of hematoxylin in the staining fluid may often be increased with advantage. 154 HISTOLOGY CHAP, PERIPHERAL NERVES AND GANGLIA. These are in general treated by the same methods as the central nervous system. Good preparations can often be obtained by teasing. Mays has described the following method for showing non- medullated nerves and nerve-endings in muscle. Mays’s Method. (1) The pieces of tissue are thoroughly soaked in a 5 per cent solution of arsenic acid (As,O,). (2) Transfer for ten minutes to the following solution :— 1 per cent solution of the double chloride of gold and potassium : 4:0 ccm. 2 per cent osmic acid é 10 ~—, 0°5 per cent arsenic acid 200 =, (3) Wash in water. (4) Transfer to a 1 per cent solution of arsenic acid, and expose to sunlight for three hours on a water-bath at a temperature of 45° C. in this solution. (5) Clear in the following mixture :— 25 per cent hydrochloric acid 1:0 ccm. Glycerine . : . 40-0 Water ‘ ‘ 20°0 ” 2 The method of injecting the tissues with methylene blue has lately been extensively used for demonstrating peripheral nerves and ganglia. The principle was first employed by Ehrlich, but has since been considerably modified. Methylene Blue Injection Method. (1) A 4 per cent methylene blue solution (preferably Dr. Griibler’s “Rectificertes methylen blau zur vitalen injection”), having been previously filtered, is injected at once, or as soon as possible after death, into the blood-vessels of the organs the nerve-structures of which are to be examined. The organ which has been thus treated is left exposed to the air until it becomes blue. This point is determined by teasing out fresh pieces of the tissue. xIV ‘EXAMINATION OF SPECIAL TISSUES AND ORGANS 155 (2) As soon as the colouration has become general, the colour is fixed by placing the tissue in a cold saturated aqueous solu- tion of ammonium picrate, which should be filtered before using. If sections are to be cut, the tissue is placed for twenty minutes to twelve hours, according to the size of the piece, in a cold satu- rated alcoholic solution of ammonium picrate. The following fixing solution is sometimes better :— Saturated aqueous solution of ammonium picrate 100 ccm. 1 per cent osmic acid : : ‘ De ss (3) Sections may be cut between pieces of lardaceous liver, elder-pith, etc. (vide p. 27), or with the freezing microtome. Sections or teased preparations should be mounted in a mixture of equal parts of glycerine and water, to which a trace of ammonium picrate is added. If other constituents of the tissue, such as cement substance, nerve cells, blood corpuscles, etc., become stained at the same time, the nervous tissues are so well defined that they are quite unmistakable. Degeneration changes in peripheral nerves can be detected by hardening in Flemming’s or in Marchi’s solutions, by which means degenerated parts of the medullary sheaths are stained black. Obersteiner has recently recommended Platner’s method with ferric chloride and dinitro-resorcin for showing degeneration in axis-cylinders. Platner’s Method. (1) Harden the nerve-trunk8 for one to several days in the following solution :— Liquor ferri sesquichloridi 1:0 com. Distilled water . é 40, (2) Wash in water until the washings cease to give a reaction with potassium sulphocyanide. (3) Transfer the pieces of nerve to a solution prepared, by adding an excess of dinitro-resorcin to 75 per cent alcohol for several weeks, according to their size. (4) Dehydrate; embed; cut. By the combination of the iron solution and dinitro-resorcin 156 HISTOLOGY CHAP. the axis-cylinders acquire an emerald-green colour, and changes in them are very clearly defined. The method can be applied to preparations which have been hardened in Flemming’s solution or in Miiller’s fluid. 1 Degeneration of peripheral nerves. I am indebted to Dr. Sherrington for the following details, which are of considerable practical importance :—When trying to determine whether any of the fibres remain normal, or whether all are degenerated, it should be borne in mind that this point cannot be ascertained by transverse sections, because, although all the fibres may be undergoing degeneration, many of them will still appear normal in cross-section, even when the degeneration is advanced (e.g. three to four weeks after section of the nerve-trunk). The degenerating fibres break up into short lengths, and in many places the transverse section of these pieces is indis- tinguishable from the transverse section of a normal fibre. It is necessary, therefore, to cut longitudinal sections, preferably a short series in paraffin, or to tease out a piece of the fresh nerve. For treating peripheral nerves °5 per cent osmic acid is better than 1 per cent, as it penetrates better, and renders them less brittle. —Ep. XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 157 THE EYE. For preparing sections of the entire eye Miiller’s fluid and Flemming’s solution are the only hardening reagents employed. Eyes can be kept for a very long time in Miiller’s fluid. Embedding in celloidin is necessary for a detailed examination of the eyeball. By this means the relative position of the layers is preserved. Staining in bulk with Bismarck brown (p. 49). or Beale’s carmine (ibid.) is sometimes advisable, so as to minimise the risk of disturbing the relation of the different parts. Embedding in paraffin may also be employed. Hematoxylin and alum carmine are used for nuclear-staining. The retina and optic nerve should be treated by the various methods already described for the central nervous system (vide p. 135). Double-staining is often useful. Cohnheim’s gold chloride method (p. 140) should be employed for the cornea. The cornea, taken from a freshly-killed animal, is placed for five minutes in fresh filtered lemon juice. It is then transferred for twenty minutes to a 1 per cent gold chloride solution, and, lastly, is exposed to the light for three to four days in water slightly acidified with acetic acid. It is embedded in celloidin and cut in alcohol. THE EAR. Portions of the middle and inner ear should be hardened in Miiller’s fluid. For the external ear alcohol may be used as well. Decalcification of the adjacent bony tissues, when necessary, is postponed until the hardening process is completed. OSSEOUS SYSTEM. Bones and joints are hardened in Miiller’s fluid. Prepara- tions hardened in alcohol do not give such good results. Decalci- fication should be done after complete hardening. Preparations hardened in Miiller’s fluid are first thoroughly washed, left in 158 HISTOLOGY CHAP. alcohol for some days, and then decalcified. (For decalcifying methods vide p. 17.) Stain with hematoxylin; double-staining with hematoxylin and neutral carmine gives very good results, as the carmine stains decalcified bones and ossifying cartilage red, but not tissue which has undergone calcareous infiltration. In this process the sections stained in hematoxylin are washed for twelve to twenty-four hours in water before placing ‘them in carmine. The ground substance of decalcified cartilage from which the lime salts have been dissolved stain deep bluish violet with hematoxylin. Unaltered cartilage varies in its behaviour to neutral carmine and hematoxylin; resting cartilage generally stains better with carmine, while, on the other hand, growing and_ proliferating cartilage stains deeper with hematoxylin. The cells of the marrow of bones are very well defined by double-staining with hematoxylin and eosine. Pommer has published a method for studying the structural arrangement of bone, and for detecting areas in bones which con- tain no lime salts. The method depends on a peculiar action of Miiller’s fluid. Bones left in Miiller’s fluid for a long time ‘become capable of being cut, owing to the feebly decalcifying action of its acid salts. In addition to this, however, it differen- _ tiates between calcified and non-caleareous portions. This is not the case with bones decalcified by acids. The method is as follows :— Pommer’s Method. The bones should be kept in Miiller’s fluid until they can be cut easily with a sharp razor. In the sections from bone thus prepared the decalcified bony matter is clearly differentiated by its homogeneous appearance from the parts which have not undergone calcification, and which are © conspicuous by their distinctly fibrillar structure. By treating with carmine the detection of small non-calcareous areas is much facilitated, and the relations of the different parts in sections become much plainer. Pommer recommends one of the following solutions for staining bones which have been decalcified in acid mixtures, such as Von Ebner’s fluid -— XIV EXAMINATION OF SPECIAL TISSUES AND ORGANS 159 (1) Dahlia ‘ . 0°04 per cent solution. (2) Saffranine . . 0°01 to 0°16 per cent solution. (3) Methyl-green . 0°30 per cent solution. The decalcified parts stain rather deeply in twelve to eighteen hours in any of these solutions, and rather more with saffranine and dahlia than with methyl-green. _ The parts which do not contain calcareous material, on the contrary, remain quite colourless, and contrast most distinctly with the parts which previously contained lime salts. Scheffer has also recommended a method for staining bony tissues with saffranine. Scheffer’s Method. (1) Decalcify with nitric acid or with an acidified sodium chloride solution. (2) Stain for a half to one hour in an aqueous solution of saffranine (1 in 2000). (3) Wash in water. (4) Transfer to a 0:1 per cent corrosive sublimate solution. (5) Examine in glycerine. Contact with alcohol should be avoided as far as possible. To prepare permanent specimens, therefore, the sections, after treatment with corrosive sublimate (4), should be rapidly passed through alcohol, and the greater part of the water removed by pressure with blotting-paper ; the rest can be got rid of by leaving them for a long time in bergamot oil. They can then be mounted in Canada balsam. MUSCLES, TENDONS, SYNOVIAL SHEATHS, AND BURSAE. These should be hardened in Miiller’s fluid. Sections of muscle cannot be prepared satisfactorily without embedding in celloidin. Picro-carmine (p. 48) is a very suitable stain for muscle; it stains the protoplasm yellow. Hematoxylin and alum carmine may also be used, or double-staining with hematoxylin and carmine or eosine. 7 Picric acid (p. 54) also is useful for double-staining. 160 HISTOLOGY CHAP. XIV MALE AND FEMALE SEXUAL ORGANS. Harden in Miiller’s fluid. Embed in celloidin, especially the ovary and placenta. ‘To examine fragments of the uterine mucous membrane for carcinoma, the pieces should be hardened in alcohol and embedded in celloidin. It is advisable to fix several pieces on one cork and cut them all at the same time. This is readily done by sur- rounding the cork with paper so as to form a box (vide p. 23), and in this way several sections from several different fragments © can be examined at once in one preparation. The examination is thus rendered much easier and more rapid, as very often the first portion, which has been cut by itself, may contain no part suitable for examination. There is also a much greater chance of cutting sections containing the muscular tissue immediately underlying the mucous membrane. Single-staining with hematoxylin is used in these cases. Shreds of decidua and remains of placenta can be examined in the same way. CHAPTER XV MICROSCOPICAL EXAMINATION FOR MEDICO-LEGAL PURPOSES Detection of Blood. In stains on wood, metal, or clothing, the examination should be directed to— (1) The recognition of red blood corpuscles. (2) Obtaining evidence of the presence of blood pigment. I. To find blood corpuscles in recent stains some of the scrapings are soaked in 0°6 per cent salt solution (distilled water should not be used), and examined directly under the microscope. The blood corpuscles of man and of mammals are circular and without nuclei, those of other animals being oval and nucleated, so that so far the examination determines whether, on the one hand, the stain is due to the blood of man or mammals, or, on the other hand, to that of some other class. / Human red blood corpuscles are larger than those of other mammals. The diameter is 0:°0077 mm. Following them in order of size come the red corpuscles of the dog, rabbit, pig, ox, horse, cat, and, lastly, the sheep. Human can be distinguished with fair accuracy from other mammalian blood corpuscles by measurement with a micrometer. As many corpuscles as possible should be measured to ensure accuracy. An absolute diagnosis is prevented by the very rapid changes in shape and form of the corpuscles. In older blood stains, red corpuscles which have retained their characteristic shape may, however, often be found. Generally, simple salt solution is not sufficient for the examination, and other fluids have to be employed. The following are the most useful :— M 162 HISTOLOGY CHAP. (1) 30 per cent caustic potash. In using this care must be taken not to dilute with water. (2) Roussin’s fluid. Glycerine ‘ é : 3 parts. Strong sulphuric acid : ‘ 1 part. (3) Pacini’s fluid (modified). Corrosive sublimate “ : 1:0 grm, Sodium chloride ‘ i 2°0 grms. Glycerine ‘ 5 . 100-0 cem. Water i : . 38000 ,, These fluids are used in the following manner :— A portion of the stain is scraped off, or, in the case of clothes or material, is removed, placed on a slide, and the fluid added to it. The blood corpuscles as they become visible are examined at once under the microscope, as they gradually undergo changes in the macerating fluid. , In old blood stains in which the corpuscles have undergone much more shrinking, differences in size of the corpuscles are much less obvious than in recent cases. Great care must be taken to avoid mistaking the spores of some of the moulds for corpuscles. Spores resist very strongly the action of acids and alkalies. II. The detection of blood pigment by the preparation of hemin crystals is a more reliable test for blood. Hemin, the hydrochlorate of hematin, is a derivative of hemoglobin. Hesemin crystals are known also as Teichmann’s blood crystals, after their first observer. They can be prepared from dried blood as follows :— If the blood stains are on a hard substance such as wood, they are carefully scraped off, and the red mass collected on a glass slide or in a watch-glass, and dissolved in a small quantity of distilled water. After removal of extraneous matter, etc., the reddish coloured fluid is evaporated down and a drop of 0°6 per cent salt solution, the size of a pin’s head, added. This is spread out on the surface of the slide and again dried. When this is done, the dried mass is scraped up with a scalpel, and glacial acetic acid added with’ t xv EXAMINATION FOR MEDICO-LEGAL PURPOSES 163 a glass rod (the test is very uncertain with dilute acid). A cover- glass is then placed on it, and it is heated over a spirit-lamp until the acetic acid begins to bubble. Gentle warming is continued until the acetic acid is evaporated. When this has taken place the brown hemin crystals separate out. Larger hemin crystals are obtained the more slowly the evaporation has been carried on. Evaporation on this account should be performed under a cover-glass. Hemin crystals are quite insoluble in water, ether, and alcohol. They are soluble with difficulty in ammonia, dilute sulphuric acid, and in nitric acid, but readily dissolve in caustic potash. They form small rhombic plates. The presence of fatty substances often hinders crystallisation ; in such cases it is advisable to treat the material beforehand with ether. The presence of rust from iron may also hinder the separation of hemin crystals, EXAMINATION OF HAIR. The first point to decide ‘in all medico-legal examinations is whether the hair in question is human or from an animal. With this in view, attention must be paid to the following microscopical features :— (1) The outermost layer of the hair, the cuticle, is composed of fine scales of epidermis lying upon one another like slates on a roof, with their pointed ends directed towards the free end of the hair. If these cells cannot be clearly made out without further treatment, dilute nitric acid should be added. In most animals the cells of the cuticle are much larger than in man, so that they are much more clearly defined; and in many animals the cuticle cells stand out more, so that they give a much more serrated appearance to the hair than is the case in man. _._. (2) The middle layer or cortical substance is composed of elongated corneous cells. These are rendered more distinct by the addition of dilute nitric acid. In human hair the cortical substance greatly exceeds in breadth the innermost layer of medullary substance. In animals the proportion is quite the reverse. (3) The cell structure of the medulla in human hair is very indistinct, while in animals it can readily be made out. In 164 HISTOLOGY CHAP, human hair the medulla is often wanting, and when present its continuity is frequently interrupted. This is rarely the case with animals, and when it occurs is always limited to quite isolated hairs. It is obvious that the hair should be examined throughout its whole length. It is advisable in each individual case, in order to decide whether the hair is human or from some animal, to compare it not only with human hair, but also with hair from such animals as are ordinarily met with. Characteristics of Hairs from Different Parts of the Body. Hairs from the beard are usually the thickest, 0-14 to 0°15 mm. in diameter; then follow in order female pubic hairs, hairs from the eyelashes, male pubic hairs, hairs from the male scalp, and, lastly, hairs from the female scalp, about 0:06 mm. in diameter. The frequency of individual variations should always be borne in mind. Differences due to age are also very considerable, the hairs of new-born children being dis- tinctly thinner than those of older children, and especially of adults. Hairs of new-born children have a pointed extremity, as have also hairs which have undergone no disturbance during their natural growth, such as cutting, pressure from clothing, softening from perspiration, etc. Cut hairs at first exhibit a sharp section, but later have a more rounded termination. Hairs which have been pulled out generally have an open, club-shaped root, with the remains of the hair sac; hairs which have dropped out have a closed, smooth, atrophied root. To decide whether the hair in question comes from a par- ticular individual, methods of comparison should be employed, attention being paid to the size of the entire hair as well as of the various layers, and as to colour, etc. For this purpose sections can be cut: with a razor; embedding in. paraffin (vide p. 24) is preferable. Detection of Seminal Stains. In examining stains for spermatozoa it should first be ascer- tained whether fine scales can be separated from the more or less firmly adherent patches on the material. Such scales, which xv EXAMINATION FOR MEDICO-LEGAL PURPOSES 165 must be very gently handled on account of their brittleness, should be mixed upon a glass slide with a drop of distilled water, and then teased up with a pair of needles. The specimen is then examined under a high power with a narrow diaphragm. If the stain has soaked deeply into the material so that scales cannot be separated from it, a small portion should be cut out and macerated for a quarter to half an hour in a watch-glass containing distilled water. By adding a small quantity of hydro- chloric acid (1 drop to 40 ccm. water), the Spermatozoa are pre- vented from swelling up. The piece of stuff is then squeezed by pressing on it with the handle of a needle, and the milky fluid thus obtained may then be examined directly. The examination can also be carried on by teasing a small piece of the material on a glass slide in water. It is very essential that the stuff should have been well soaked before- hand. , If spermatozoa are found, cover-glass preparations may be obtained in the ordinary way, and stained with neutral carmine or eosine. Hematoxylin and eosine are used for double- staining. Unger recommends a special staining solution :— Methyl-green ‘ . 0°15 to 0°3 grms, Water. ; . 100°0 ccm. Hydrochloric acid 3 drops. This solution may be used for soaking the piece of material under examination, and it is then examined directly under the microscope. It may also be employed for staining the cover-glass preparations. The hinder part of the head stains dark green, the anterior portion light green, while the median part and the tail stain a lighter colour than the hinder part of the head. EXAMINATION OF DECIDUAL REMAINS. This may be necessary in cases where there is suspicion of induced abortion. The decidua of pregnancy can be distinguished from all other constituents of tissues by the characteristic large polygonal or rounded cells. 166 HISTOLOGY CHAP. XV Fresh teased preparations may be made, but it is better to cut small sections after hardening in alcohol. These can be cut either by hand or in a cylinder microtome (vide p. 32). After surrounding with paraffin, they can also be cut between fresh elder-pith or hardened liver (wide p. 27). They stain best with hematoxylin. INDEX Assi, illumination apparatus, 2 Acarus folliculorum, 108 scabiei, 108 Acetic acid, for fresh tissues, 10 Acid, acetic, 10 chromic, 17, 58 hydrochloric, 11, 19 nitric, 20 osmic, 15 picric, 15, 19 Acid-alcohol, 47 Actinomyces, 103 Adamkiewicz’s saffranine stain, 148 Albumen, cement for paraffin sections, 26 Alcohol, absolute, 12 acid, 47 hardening in, 12 for showing karyokinesis, 59 Alimentary canal, 129 Altmann, fixing method, 59 cell inclusions, 60 Alum carmine, 46 Alum hematoxylin, 44 Ammonium phosphate, 133 sulphide, 70 Amyloid disease (v. lardaceous), 65 Anchylostomum duodenale, 108 Aniline acetic acid, 78 blue, 139 oil, 78 water, 78 water saffranine, 57 Anthrax bacillus, 92 Apathy, hematoxylin method, 46 Apochromatic lenses, 1 Apparatus for histological work, 1 Arnold’s method, 8, 73 Aronson-Philipp’s stain for granules, 118 Arteries, 124 Ascarus lumbricoides and vermicularis, 108 Aspergillus, 107 Asphalt varnish, 42 . Asthma crystals, 1384 Atrophy, 63 . Axis-cylinder staining, 137 et seq. BaBEs, aniline-water saffranine, 57 Bacilli, methods of staining, 81 Bacillus, anthrax, 92 charbon syptomatique, 94 cholera, 102 diphtheria, 92 Friedlinder’s encapsuled, 91 glanders, 94 lepra, 101 mouse septicemia, 96 rhino-scleroma, 93 swine erysipelas, 96 syphilis, 97 tetanus, 94 tubercle, 97 typhoid, 96 Bacteria, methods of staining, 77 effects of Gram’s method upon, 81 epiphytic, 128 flagella of, 81 in blood, 122 in fluids, 76 in sections, 83 spores of, 87 Balzer, stain for moulds, 107 Baranski, stain for actinomyces, 105 Baumgarten, method for karyokinesis, 60 stain for leprosy bacillus, 101 Beale’s carmine, 49 Benda’s hematoxylin, 58 Bergamot oil, 43 Berlin blue, 29 Biedert’s sediment method, 98 Bignani, stain for plasmodia, 109 Bile capillaries, 131 Bilirubin, 133 Biondi, examination of blood, 119 and Heidenhain’s stain, 51 Birch-Hirschfeld, stain for lardacein, 67 Bismarck brown, 49 Bizzozero, stain for bacteria in skin, 128 and Vassale’s method, 59 Blood, bacteria in, 122 counting corpuscles, 123 cover-glass preparations, 113 examination of, 111 pigment, 122 platelets, 120 preservation of, 112 sections of, 119 168 “HISTOLOGY Blood stains, 161 Boeck, bacteria in skin, 128 Boehm, method for bile capillaries, 131 ' stroma of liver, 131 Boiling, method of, 16 Bone, examination of, 157 decalcification of, 17 methods of staining, 158 Borax carmine, 47 Bostroem, stain for actinomyces, 104 Bousfield, staining flagella, 82 Brushing sections, 59 Burse, 159 Catcrum, carbonate and phosphate, 68, 133 Caldwell, celloidin embedding, 23 Camera lucida, 4 Canada balsam, 42 Carbol-fuchsine, 78 Carbolic acid water, 78 Carmine, alum, 46 Beale’s, 47 borax, 47 lithium, 47 neutral, 52 picro-lithium, 49 Casts, 133 Caustic potash, 8 Cedar-wood oil, 43 Cell inclusions, 60 Celloidin embedding, 22 and paraffin method, 26 Cellulose, 66 Cements for cover-glass, 42 Central nervous system, 135 Centrifugalisation, 9 Cholera, bacillus of, 102 detection in intestine, 130 Chromic acid for decalcification, 17 dissociating, 8 karyokinesis, 58 Chrom-formic acid, Rabl’s, 58 Chrom-picric acid, 59 Ciaglinski, axis-cylinder stain, 139 Clearing reagents, 43 Cloudy swelling, 63 Clove oil, 43 Coccidia, 108 Cochineal-alum, 138 Cohnheim, gold chloride method, 140 injection fluid, 29 Collodionised plates, 37 Colloid material, 65 Condenser, 2 Copper sulphate, for dehydration, 12 Cornea, 157 Corpora amylacea, 66 Corpuscles of blood, size of, 161 Corrosive sublimate, hardening, 14 for nerve-cells, 149 karyokinesis, 58 Counting corpuscles, 123 Cover-glasses, 4 Cover-glass preparations, 79 Curschmann’s spirals, 134 Cutting in series, 36 Cylinder microtome, 32 Cystin, 133 Ozaplewski stain for tubercle, 101 Czokor, cochineal-alum stain, 138 Damar varnish, 41 Darkschewitsch, cutting in series, 38 Decalcifying methods, 1 Decidual remains, 160, 165 Decolourising reagents, 78 Degeneration, colloid, 65 fatty, 63 hyaline, 68 in peripheral nerves, 156 lardaceous, 65 mucoid, 64 Dehydration, 42 Delépine, interlamellar films, 76 Digestion, artificial, 38 Diphtheria, bacillus of, 92 Diplococcus pneumoniz, 90 Dissociating or macerating fluids, 8 Distoma hepaticum, 108 Double-staining, 40, 52, 68 Doutrelepont, stain for syphilis bacillus, 97 Drawing apparatus, 4 Dry method for sections, 87 Ear, examination of, 157 Ebner, decalcifying fluid, 19 Echinococcus, 110 Ehbrlich-Biondi method, 51 Ehrlich, acid-hematoxylin, 45 cover-glass method, 113 examination of blood, 114 gentian violet solution, 59 hematoxylin-eosine solution, 54 “ mastzellen,” 73 tubercle stain, 99 Elastic fibres in sputum, 134 in tissues, 126 Elder-pith, 27 Embedding methods, 22 Endocarditis, 125 Fosine solutions, 53 Eosinophile cells, 114 Erlicki, hardening fluid, 14 Exner, stain for medullated nerves, 147 Eye, examination of, 157 Eye-pieces, 1 Farrants’ medium, 42, 49 Fat, reactions of, 63 Fatty degeneration, 63 Fibres, elastic, in skin, 126 in sputum, 134 Fibrin, staining methods, 121 Flagella, methods of staining, 81 Flechsig-Golgi stain for nerve-cells, 151 Flemming’s solution, 15, 56, 64 Flormann, stain for actinomyces, 106 Fluids, examination of, 9 Fluorescin, 79 Foa, fixing solution, 59 INDEX 169 Fol, decalcifying fluid, 18 fixing solution, 59 Fraenkel, C., examination of moulds, 107 Freezing microtome, 34 Fresh tissues, examination of, 7 sections of, 27 staining, 11, 49 Freud, gold chloride method, 140 Friedlinder, encapsuled bacillus, 91 Fuchsine, 84 Fuchsine-acetic acid, 10 Fungi, 107 GaBBET, tubercle stain, 100 Ganglion cells, 136 Gelatine cultures, sections of, 87 Gentian violet, Ehrlich’s solution, 59 nuclear stain, 50 stain for bacteria, 84 Giacomi, stain for syphilis bacillus, 97 Giesson, van, method of staining, 54, 65, 68 Glanders, bacillus of, 94 Glass apparatus, 5 Glycerine, for fresh tissues, 9 mounting in, 41 Glycerine-gelatine cement, 33 Glycogen, 68 Gold chloride staining, Cohnheim’s, 140 Freud’s, 140 Upson’s, 141 é Golgi, silver stain for nerve-cells, 149 Golgi-Flechsig, stain for nerve-cell pro- cesses, 151 Gonococcus, 89 Gram, stain for bacteria, 80, 84 method for karyokinesis, 60 Granules in leucocytes, 115 Gudden’s microtome, 34 mixture, 34 Gum and celloidin plates, 36 Gum-glycerine, 15 Gum solution for freezing, 35 Giinther, decolourising method, 80 H=MATEIN or hem-alum, 45 Hematocytometer, 123 Hematoidin, 69 Hematoxylin staining, 45 Apathy’s method, 46 Benda’s method, 58 Ebrlich’s method, 45 Heidenhain’s method, 46 Kulschitzky’s method, 146 Weigert’s method, 142, 144 Hematoxylin and eosine, 68, 54 carmine, 54 Hemosiderine, 69 Hanging-drop method, 76 Hardening methods, 12 Haug, decalcifying method, 18 Heart, 124 Heidenhain-Biondi method, 51 Hermann, fixing solution, 58 Herxheimer, stain for elastic fibres, 126 Honegger, neutral carmine, 52 Hoyer, stain for mucin, 65 Hyaline degeneration, 68 Hydatid cysts, 110 Hydrochloric acid, 11 for decalcifying, 19 IMMERSION lenses, 1 Impregnation with lime salts, 69 Inflammation, 74 Injection methods, 28 fluids, 29 Injection of lymphatics, 30 Interlamellar films, 76 Intestinal contents, 130 Iodine, solution of, 10 green, reaction with lardacein, 67 reactions, 65, 66, 68 sulphuric acid, 66 Tris diaphragm, 3 Tron, detection of, 69 Israel, stain for actinomyces, 105 KaryoxinEsis, 51, 56 Kidney, 182 Koch, stain for tubercle, 98 comma bacillus, 130 Kollmann’s injection fluid, 30 Kiihne, methylene blue method, 86 Kulschitzky, stain for nerve-fibres, 146, 147 Lactic acid for decalcifying, 19 Leevulose as a mounting medium, 67 Lamps, 3 Langhans, detection of glycogen, 68 Lardacein, 65 Lardaceous degeneration, 65 Lardaceous liver for embedding, 27 Lavender oil, 43 Leonhardi’s ink, 67 Leprosy bacillus, 101 Leucin, 133 Leucocytes, 113 Lifter, 5 Lime salts in tissues, 68 Lithium carmine, 47 Liver, 130 Léffier’s methylene blue, 79 stain for bacteria, 84 stain for flagella, 81, 89 Logwood, vide hematoxylin, 45 Lowit, stain for fibrin, 122 Lugol’s solution, 10 Lung, 133 Lustgarten, bacillus of syphilis, 97 Lymphatic glands, 125 Lymphatics, injection of, 30 Lymphocytes, 114 MacEratIne or dissociating fluids, 8 Malaria, plasmodia of, 109 Malbory, axis-cylinder stain, 153 Manchot, stain for elastic fibres, 127 Marchi’s method, 135 Mastzellen, 73, 81, 122 Mays, stain for nerve-endings, 154 170 HISTOLOGY Methylene blue, Kiihne’s solution, 86 ‘Liffler’s solution, 79 injection method, 154 use in photography, 61° Methyl green, 67 Methyl violet, 67 Micrococci, 89 Micrometer eye-piece, 4 Microscope, 1 Microtome, cylinder, 82 freezing, 34 Gudden’s, 34 sliding, 33 Moller, staining spores, 88 “Moulds, 107 Mounting fiuids, 41 Mucoid degeneration, 64 Mucous membranes, 129 Miiller, H. F., stain for blood, 120 Miiller’s fluid, 8, 12 Muscle, 159 Muscle, trichinae in, 108 Naputua, 48 Necrosis, 62 Neisser, sections of gelatine cultures, 87 Nerve- cells, 136 Nerve-cell processes, 149 Nerves, peripheral, 154, 156 Nervous system, central, 135 Neutral carmine, 52 Nicolle and Morax, stain for flagella, 82 Nikiforoff, stain for nervous tissues, 148 blood, 114 nuclear-staining, 52 stain for spirilla, 102 Nissl, stain for nerve-cells, 136 Nitric acid, 20 Noniewicz, stain for glanders bacillus, 95 Nuclear-staining, 44 Nuclear division, 51, 56 OBJECTIVES, 1 Obregia’s method, 150 Oil immersion lenses, 1 Oppel, stain for fibrous tissue in liver, 131 Orcein for actinomyces, 105 Origanum oil, 43 Orseille, for actinomyces, 105 Osmic acid, for hardening, 15 decalcifying, 18 fat, 11 dissociating, 8 “ fixing,” 56 Osseous system, 157 Pacini’s solution, 112, 162 Pal-Weigert method, 145 Palladium chloride, 20 Pancreas, 130 Paraffin, embedding in, 25 Paraffin and celloidin method, 26 Parasites, 108 ' Peripheral nerves and ganglia, 154, 156 Pfeiffer’s fuchsine method, 84 Phloroglucin, ‘for decalcifying, 20 Phosphates, detection of, 72, 133 Picric acid, 15, 19, 54 Picro-carmine, 48, 49 Picro-lithium carmine, 49 Pigment, 69, 134 Pigmentary atrophy, 63 Placenta, 160 Plasmodia in blood, 109 Platner, ferric chloride method, 155 Plehn, stain for plasmodia, 109 Pneumonococcus, 90 Pommer, stain for bone, 158 Potash, caustic, 8, 11, 78 Potassium acetate, 10, 42 ferrocyanide, 69, 71 Protozoa, 108 Pyogenes aureus, 81, 89 Pyroligneous acid, 19 QuincEE, detection of iron, 72 RaBL, chrom-formic acid, 58 Ramon-y-Cajal, stain for nerve-cells, 150 Rauschbrand bacillus, 94 ; Ray fungus or actinomyces, 103 Razor, 31. Reagents for fresh tissues, 8 Recurrent fever, spirillum of, 102 Respiratory organs, 133 Revolving nose-piece, 4 Rhino-scleroma, bacillus of, 93 Ribbert, stain for Friedlinder’s bacillus, 92 Rindfleisch, method for blood, 120 Roussin’s fluid, 162 Russell, cell-inclusions, 61 SaFFRANINE, 57, 148 Sahli, stain for nerve-fibres, 139 Saline solution, 7, 112 Sarcina ventriculi, 92 Scheffer, stain for bone, 159 Schmaus, axis-cylinder stain, 138 Schottelius, cholera bacilli in intestine, 130 Schutz, stain for gonococci, 90 glanders bacillus, 95 Sections, 31 artificial digestion of, 38 brushing, 38 dehydrating, 42 mounting, 41 of blood, 119 of gelatine cultures, 87 shaking, 38 staining, 39 Sediment method, 98 Septiceemia bacillus of mouse, 96 Serial sections, 36 Serous fluids, 125 Serum, artificial, 7 Sexual organs, 160 Shellac solution, 26 ‘*Siebdose,”” Steinach’s, 5 Silver nitrate method, 149 Skin, 125 INDEX Spermatozoa staining, 165 Spirillum of recurrent fever, 102 - Spleen, 125 Spore-staining, 87 Sputum, 98, 133 Staining, general methods of, 40 Staining, bacteria, 77 diffuse, 52° double, 40, 52 fresh tissues, 49 in bulk, 55 karyokinetic figures, 56 nuclear, 44 spores, 87 Staphylococcus pyogenes, 125 Starch, 66 Stepanow, stain for bacillus of rhino- scleroma, 93 Stieda, detection of iron, 70 Strasser, cutting in series, 37 Streptococci, 125 Stroebe, axis-cylinder stain, 139 Stroschein, sediment method, 99 Sublimate, corrosive, for hardening, 14 for karyokinesis, 58 for nerve-cell processes, 151 Syphilis, bacillus of, 97 TaEnia, 110 Taenzer, stain for elastic fibres, 127 Teasing methods, 7 Teichmann’s crystals, 162 Tendons, 159 Tetanus bacillus, 94 Thiersch, injection mass, 3 Thionin, 65 ‘ Thoma, decalcification method, 21 Thoma-Zeiss apparatus, 123 Tissues, fresh, 7 hardening, 12 staining, 40 Toison’s diluting fluid, 123 Toluidine blue, 65 Tongue, ulcers of, 9 Touton, stain for gonococci, 90 Trichina, 108 Triple phosphates, 133 Tropeolin, 79 THE 171 Trypsin solution, 39 Tubercle bacilli, 97 _ in urine, 182 Tumours, 73 Turpentine, 25 Typhoid bacillus, 96 Tyrosin, 133 Uucers of tongue, 9 Unger, stain for spermatozoa, 165 Unna, bacteria of skin, 129 dry method for sections, 87 elastic fibres, 126 Unna-Taenzer, stain for skin, 127 Upson, stain for axis-cylinders, 141 Urates, 133 Uric acid, 133 Uterus, 165 Van GrEsson, method of staining, 54, 65, 68, 137 Van Ketel, tubercle bacilli in urine, 132 Vassale’s method, 145 Vegetations, 125 Vesuvine or Bismarck brown, 49 Von Ebner’s fluid, 19 WALDEYER, decalcifying method, 18, 20 Water-immersion lens, 1 Weigert, cutting in series, 36 picro-carmine, 48 stain for actinomyces, 105 bacteria, 86 fibrin, 62, 121 white matter, 142, 144 Westphal, stain for ‘‘mastzellen,” 74, 117 Wolkowitsch, staining bacillus, 93 Wolter’s axis-cylinder stain, 152 rhino - scleroma XyYLOL, 25, 43 Canada balsam, 42 carbolic acid, 43 ZALEWSKI, detection of iron, 70 Ziehl-Nielsen, tubercle stain, 100 END Printed by R. & R. Ciark, Edinburgh Hee ea eee Rua reer a oe Sous oa reas oe ier Escer i s eR