A PRACTICAL MANUAL OF MEDICAL AND BIOLOGICAL STAINING TECHNIQUES «3 fcr" 7^ A PRACTICAL MANUAL OF MEDICAL AND BIOLOGICAL STAINING TECHNIQUES hy EDWARD GURR F.R.I.C., F.R.M.S., F.L.S., M.I. Biol. INTERSCIENCE PUBLISHERS, INC. NEW YORK FIRST EDITION, 1 95 3 SECOND EDITION, 1 956 PRINTED IN GREAT BRITAIN BY J. W. ARROWSMITH LTD., BRISTOL PREFACE TO FIRST EDITION This book has been written because it is felt that there is an urgent need of a practical manual dealing with all or most branches of microscopic staining, entirely divorced from theory and general statements. It has been the writer's experience that a great deal of time is lost in searching through volumes of theory and general statements in order to extract a particular staining method which, even when found, may not be complete. It is hoped that this book will form a useful supplement to the standard works on anatomy, bacteriology, biology, botany, cytology, embryology, entomology, histology, mycology, path- ology, veterinary science, zoology, etc. It should be pointed out that this book contains all the subject matter, revised, rearranged and co-ordinated, of the writer's own publications Microscopic Staining Techniques which were put out in the form of three booklets. No. i, 2 and 3 as a temporary meas- ure in an endeavour to meet the more pressing demands for infor- mation on the application of microscopic stains. The booklets attained a world-wide circulation and were in fact chosen by the British Council for inclusion in their exhibits of British medical books. The writer has since received many requests to incorporate all three parts of Microscopic Staining Techniques into one and this has been done in the present book, but with many additions. Many of the methods given here are standard, some are not so well known, while others are unknown. In writing this book, references have been made to numerous journals and standard works, chief of which were as follows : Stain Technology (Biotech Publication, Geneva, N.Y., U.S.A.). Plant Microtechnique, by D. A. Johansen (McGraw-Hill, New York). Handbook of Practical Bacteriology, by T. J. Mackie and J. E. McCartney (E. & S. Livingstone, Edinburgh). McClung's Handbook of Microscopical Technique, edited by Ruth McClung Jones (Paul B. Hoebber,Inc., New York, 16, U.S.A.). PREFACE Laboratory Tech?iique, by E. V. Cowdry (Williams & Wilkinson, Baltimore, U.S.A.). Histopathologic Technic, by R. D. Lillie (Blakiston Co., Phila- delphia, U.S.A.). Schaffer's Essentials of Histology, by H. M. Carleton and E. H. Leach (Longmans, Green & Co., London). Pathological Technique^ by F. B. Mallory (Saunders, Phila- delphia, U.S.A.). Histological Technique, by H. M. Carleton and E. H. Leach (Oxford University Press, England). The section on Fluorescence Microscopy is a modification of the writer's paper which was published in The Journal of the Royal Naval Medical Service, 1951, Vol. xxxvil, No. 3, and thanks are due to the editor of that journal for permission to in- clude the modified paper in this book. I should like to place on record my thanks to my wife, F. P. Gurr, B.Sc, for her helpful criticism of the manuscript. August, 1952 VI PREFACE TO THE SECOND EDITION I am very happy that the first edition of this book has met with so kindly a reception; and I hope that this enlarged and revised edition will be of greater service to medical research workers and biologists all over the world. Many additions have been made to the subject-matter of the book, and I wish to thank readers of the first edition for their helpful suggestions, most of which have been incorporated in this second edition. A section of histochemical methods has been added as it is felt that this may be of service to various laboratory workers, including those engaged in cancer research, who may be interested in the rapidly expanding science of histochemistry, which offers a rich field for investigation. The subject-matter of the book, as before, has been divided into various headings: some of the methods which might have been classified as " Cytological " have been placed under other headings because the particular methods are more frequently required by workers other than cytologists. For similar reasons, methods which might have been transferred to the histochemical section have been left under the same headings where they ap- peared in the first edition. In addition to the standard works mentioned in the preface to the first edition, reference has also been made to The Micro- scopisfs Vade-Mecum edited by J. Bronte Gatenby and H. W. Beams (J. & A. Churchill, Ltd., London), and Microscopic Histo- chemistry by George Gomori (University of Chicago Press, U.S.A.). It was stated in a review in a Balkan journal, and correctly so, that the first edition of this book gave no information on the tech- niques used in eastern and south-eastern European countries. The reason for this is the scarcity here of the medical and bio- logical literature of those countries. I am not a linguist, but I have painfully translated some hundreds of pages of French and Ger- man literature to find some missing link for inclusion in this present edition and, if any medical or biological laboratory workers vii PREFACE in any country in the world have any new or improved techniques that would prove to be of help to their fellow laboratory workers in other countries, I should be very pleased to be given the oppor- tunity of considering such techniques for inclusion in the third edition of this work: I am most anxious that it should be of the greatest possible service to medical and laboratory workers throughout the world and information in any language concerning particular techniques will be welcomed. It has been stated in a Netherlands journal in a review, of the first edition of this book, that Gram's Iodine and Lugol's Iodine are identical ; I must correct this misleading statement which was evidently made in error: Gram's Iodine is not identical with Lugol's Iodine; the two formulae are given in this book. I should like to express my appreciation of the very energetic co-operation afforded me by my publishers, and the efficient manner in which their production manager, Mr. R. G. Thixton, and the printers Messrs. J. W. Arrowsmith Ltd., have handled the complicated business of getting the second edition into print. 42, Upper Richmond Road West Edward Gurr East Sheen London. April, 1955 vui • Ji ' ' »Ti i CONTENTS SECTION I— GENERAL METHODS FIXATION (a) FIXATIVES Acetic Acid — Acetone — Alcohol Absolute — Allen's Fixative — Bouin's Fluid — Bouin-Duboscq (Duboscq- Brasil or Alcoholic Bouin) — Carnoy's Fluid — Carnoy- LeBrun — Champy's Fluid — Flemming's Fluids — Formalin (neutral) — Formalin (buffered) — Formol- Alcohol — Helly's Fluid — Hermann's Fluid — Klein- enberg's Fluid — Lewitsky's Fluid — Marchi's Fluid — Navashin's Fluid — Orth's Fluid — Petrunkevitch's Cupric p-Nitrophenol — Regaud's Fluid — Schau- dinn's Fluid — Susa (Heidenhain) — Zenker's Fluid, etc. (b) DEHYDRATION, CLEARING, EMBEDDING, SECTIONING, ETC Celloidin Method Celloidin - Paraffin Wax (double embedding) Celloidin - Pyridin Frozen Sections Gelatine Embedding Low Viscosity Nitrocellulose (L.V.N.) Method Page Mercuric Chloride Precipitates in Sections : Removal of 28 Paraffin Wax Embedding and Sectioning Paraffin Wax - Pyridin Method Water Wax . . . . . . Miscellaneous Dehydrating and Clearing Reagents Solvents, etc. Butyl Alcoholy Tertiary — for dehydrating and clearing Cajeput Oil — for clearing in wet climates IX 17 17 20 22 23 24 26 29 33 33 34 34 35 CONTENTS Page Miscellaneous Dehydrating and Clearing Reagents ; Solvents, etc. {continued) Cellosohe — for dehydrating . . . . . . . . 35 Dioxane — for dehydrating . . . . . . . . 36 Ethylene and Propylene Glycols — as solvents and for dehydrating . . . . . . . . . . . . 37 Propyl Alcohol — for dehydrating and clearing . . 38 Terpineol — for dehydrating . . . . . . . . 38 Mounting Media . . . . . . . . . . . . 39 Aqueous 39 Apathy Gum Syrup Mountant — Aquamount — Ber- leze Fluid — Doetschman's Gum Chloral Mountant & Stain — Glycerin — Glycerin Jelly — Glychrogel Mount- ant. Non-aqueous 41 Canada balsams — Clearmount — Cristalite — D.P.X. (Lendrum and Kirkpatrick) — Emexel — Meedol Balsam — Fluormount — Michrome Mountant, S.Q.D. Balsam — ^Venetian Turpentine SECTION 2— ANIMAL HISTOLOGY Normal and Pathological Alcian Blue - Chlorantine Fast Red forj mucopolysac- charides, connective tissue, cartilage and bone . . . . 47 Aldehyde Fuchsin (G. Gomori), for elastic tissue, mast cells, beta cells of the pancreatic islets . . . . . . 48 Aldehyde Fuchsin - Haematoxylin - Light Green - Orange G - Chromotrope for differentiating two types of basophils in the Adenohypophysis of the rat and mouse . . . . . . . . . . . . . . $0 Alizarin Red, S, for calcium deposits in cartilagenous and embryonic bone . . . . . . . . . . . . 53 Alizarin Red, S, for bone in small vertebrates (Dawson's method) . . . . . . . . . . . . . . 53 Alizarin Red, S (William's method), for mammalian embryos ; mature specimens of Urodele Amphibians ; for distinguishing between bone and cartilage . . . . 54 Alizarin Red, S, for minute bones and foetal ossification . . 56 Alizarin Red, S, for nervous tissues (Benda's method) . . 57 CONTENTS Page Alizarin Red, S, for vital staining of nervous tissue in small vertebrates . . . . . . . . . . . . 58 Alum Carmine - Aniline Blue - Orange G for demonstrat- ing the various components of the Hypophysis . . 59 Ammoniacal Silver Carbonate for vascular reticulum, tumour cells, and connective tissue around tumour in abnormal brain . . . . . . . . . . . . 60 Aniline Blue - Acid Fuchsin for elementary bodies, in sections . . . . . . . . . . . . . . 61 Aniline Crystal Violet - Gram's Iodine for epithelial nores .« .• •• .. •• •• •• •• vj^ Aniline Crystal Violet - Lithium Carmine - Iodine for fibrin and for Gram-positive organisms in tissues . . 63 Aniline Blue - Orange G (Mallory) for collagenous and reticulum fibrils ; cartilage, bone, amyloid, nuclei, fibroglia and elastin fibres . . . . . . . . 64 AzAN Stain (Heidenhain) for mucin, neuroglia, chromatin, muscle tissue, etc. . . . . . . . . . . 65 Azo Carmine - Mallory Stain for islets of Langerhans 66 Azocarmine - Haematoxylin - Acid Green - Orange G, for differential cell analysis of rat anterior hypophysis 68 Bauer - Feulgen, for glycogen, etc. . . . . . . 70 BiEBRiCH Scarlet - Ethyl Violet - Haematoxylin, for , pepsinogen granules of the body chief cells in the gastric glands . . . . . . . . . . . . . . 7^ Bismark Brown - Methyl Green for mucin, cartilage, goblet cells in embryonic tissue, trachea and intestine 72 BiONDi - Ehrlich - Heidenhain Stain for nucleoli, chrom- atin, mucin, etc. . . . . . . . . . . . . 73 Best's Carmine for glycogen . . . . . . . . . . 73 Benzidine for brain capillaries . . . . . . . . 75 Basic Fuchsin - Methylene Blue for Negri bodies in sec- tions . . . . . . . . . . . . . . 7^ Basic Fuchsin - Gentian Violet - Iodine for bacteria in oeotions .. .. .. .. ». •• •• // Carbol Aniline Fuchsin for Negri bodies . . . . 79 Carbol Fuchsin (Ziehl Neelsen) for Nissl bodies . . . . 79 Carbol Fuchsin - Borrel Blue for leprosy and for tub- ercle bacilli . . . . . . . . . . . . 80 xi CONTENTS Page Carbol Fuchsin - Haematoxylin for tubercle bacilli in mammalian tissues . . . . . . . . . . 8i Carbol Fuchsin - Haematoxylin - Picro Acid Fuchsin for M. leprae in sections . . .-. .. .. .. 82 Carbol Fuchsin - Iodine - Haematoxylin - Orange G for demonstrating leprosy organisms together with neurokeratin of the myelin sheath . . . . . . 83 Carbol Fuchsin - Methyl Green for demonstrating hyaline substance . . . . . . . . . . . . 85 Carbol Thionin - Picric Acid (Schmorl) for bone cana- i.1 w LXXX •• •• •• •• «• •• •• ^ ^ Carmine - Methylene Blue (Schultz-Schmitz Stain) for demonstrating sodium urate in animal tissues . . 86 Celestin Blue - Chromotrope 2R (Lendrum), a substitute for haematoxylin-eosin for simple diagnostic or photo- graphic purposes . . . . . . . . . . - . . 88 Celestin Blue - Orcein - Light Green (Lendrum) for breast carcinoma and skin lesions . . . . . . 89 Chlorazol Black, a general-purpose stain for whole tissues and for sections . . . . . . . . . . . . 90 Congo Red for amyloid . . . . . . . . . . 91 Congo Red - Aniline Blue - Orange G for elastic UDlCd •• •• •• •• •• •• •• V Congo Red - Ehrlich Haematoxylin for eleidin and ker- atohyalin . . . . . . . . . . . . . . 93 Cresylfast Violet - Toluidine Blue - Thionin, a tribasic stain for nerve cells and Nissl granules in normal and pathological tissues . . . . . . . . . . 93 Dahlia, Acetic (Ehrlich) for mast cell granules in sec- tions . . . . . . . . . . . . . . 95 DoPA Reagent for melanoblasts . . . . . . . . 95 Elastin Stain (Weigert) for elastic tissues . . . . 96 Elastin Stain (Sheridan) for elastic tissues . . . . 97 Elastin - Trichrome Stain for elastic, smooth muscle and collagenic fibres . . . . . . . . . . . . 97 Eosin Azur 2 - Haematoxylin (Maximow) for demonstrat- ing changes of haemopoietic tissues . . . . . . 99 Ethyl Violet - Biebrich Scarlet (Bowie) for pepsinogen granules in gastric mucosa . . . . . . . . 100 xii CONTENTS Page Feulgen Fuchsin for chromatin in animal cells . . . . loo FoNTANA Stain for argentaffine granules . . . . . . 102 FoNTANA Stain - Silver Nitrate for reticular and collagen fibres . . . . . . . . . . . . . . 102 Gallocyanin - Orcein - Acid Alizarin Blue - Alizarin Viridine, a general stain for animal tissues . . . . 104 GiEMSA Stain for malaria parasites, rickettsia, etc. . . 105 GiEMSA - Wright Stain, a permanent stain for differentiat- ing the structures, particularly Nissl bodies and cytons, of the spinal cord . . .. .. .. .. .. 106 Gold Chloride - Sublimate (Cajal) for neuroglia fibres ; for astrocytes in the central nervous system . . . . 107 GoLGi Method (Rapid) for nerve cells . . . . . . 108 Gram's Iodine for bacteria in sections . . . . . . 108 Haemalum - EosiN for demonstrating collagenous tissues 109 Haematoxylin - AzoPHLOXiN, for muscle, connective tissue, ganglion cells, etc. . . . . . . . . no Haematoxylin - Basic Fuchsin for haemofuscin, melanin and haemosiderin .. .. .. .. .. ..112 Haematoxylin (Delafield) - Eosin for general staining . . 112 Haematoxylin (Ehlrich) for keratohyalin .. .. .. 113 Haematoxylin (Ehrlich) for sodium urate in tissues .. 114 Haematoxylin (Ehrlich) - Eosin for general staining . . 114 Haematoxylin - Fluorchrome (Kultschitzky-Pal method) for myelin sheaths, etc. .. .. .. .. ..115 Haematoxylin - Gentian Violet - Iodine for Gram- positive bacteria and fibrin in sections . . . . . . 116 Haematoxylin (Heidenhain) for nuclei and cytoplasmic structures .. .. .. .. .. ..117 Haematoxylin for identification of lipines .. .. 118 Haematoxylin (Kultschitzky), Weigert's Modification for the finer studies of corticaLarchitecture and for total brain sections .. .. .. .. .. ..119 Haematoxylin - Phloxine - Aniline Gentian Violet for actinomyces in sections . . . . . . . . , . 120 Haematoxylin, Phosphotungstic (Mallory) for pleuro- pneumonia organisms in sections of lung . . . . 121 Haematoxylin - Picro Fuchsin, for nuclei, connective tissue, etc. . . . . . . . . . . . . 122 • • • XI 11 CONTENTS Page Haematoxylin - PiCRO Ponceau S, a selective stain for collagen and connective tissue in place of Haematoxylin - Van Gieson . . . . . . . . . . . . 123 Haematoxylin (Weigert) - Ponceau Fuchsin (Curtis) for collagen and connective tissue . . . . . . . . 123 Haematoxylin - Ponceau Fuchsin - Fast Green, FCF, a differential stain for pathological tissues . . . . 124 Haematoxylin - Ponceau S - Picro Aniline Blue, a differential stain for muscle and connective tissues . . 125 Haematoxylin (Weigert) - Scarlet R for demonstrating fatty acid crystals, soaps and neutral fats in fat necrosis . . . . . . . . . . . . . . 126 Haematoxylin (Ehrlich) - Van Gieson, a selective stain for collagen and connective tissue . . . . . . . . 127 Haematoxylin (Heidenhain) - Van Gieson, a selective stain for collagen and connective tissue . . . . . . 128 Hickson's Purple, a general stain for class work . . 130 Hickson's Purple - Victoria Green, G, a general stain suitable for class work . . . . . . . . . . 130 Hitchcock and Ehrlich's Mixture for lymphatic, ganglion, plasma and basophilic cells; immature cells of bone marrow, striated muscle and fibrin .. .. .. 131 Jenner Stain for blood-forming organs . . . . . . 132 Jenner - GiEMSA Stain for the polychromatic staining of blood-forming organs .. .. .. .. ..133 Lead Haematoxylin - Acid Fuchsin (MacConaill), a "definitive" polychrome stain for the central nervous system and the trunks outside it . . . . . . 134 Leishman Stain for the general differentiation of blood corpuscles; for malarial parasites ; trypanosomes, etc. 135 Leuco Patent Blue for haemoglobin . . . . . . 136 Levaditi's Stain for Treponema palHdum in sections . . 138 Light Green - Acid Fuchsin (Alzheimer) for demonstrat- ing neuroglia changes .. .. .. .. .,138 LiGNiN Pink, for whole mounts of marine vertebrates, etc. and for chitin . . . . . . . . . . . . 140 Lithium Silver (Laidlaw) for staining skin and tumours . . 141 LoRRAiN - Smith - Dietrich Stain for lipoids . . . . 143 Lugol's Iodine for the identification of glycogen . . 143 xiv CONTENTS Page LuxoL Fast Blue - Cresyl Fast Violet (Kluver & Barrera) for the combined staining of cells and fibres in the nervous system . . . . . . . . . . . . 144 MacCallum's Stain for influenza and Gram-positive organisms in tissues . . . . . . . . . . 147 Mallory Stain - Haematoxylin, for differential staining of insulin cells of the pancreas . . . . . . . . 148 Mallory Stain - Haematoxylin for differential staining of acidophils, basophils and chromophobes in mouse pituitary . . . . . . . . . . . . . . 150 Mallory Stain - Haematoxylin, for Negri bodies in sections of brain.. .. .. .. .. .. 152 Mallory Heidenhain Stain (Jane E. Cason's modifi- cation), a rapid one-step method for connective tissue 153 Mallory's Phosphotungstic Acid Haematoxylin, a general stain for vertebrate tissues . . . . . . 154 Marshall Red - Victoria Green, a general stain for class work . . . . . . . . . . . . . . 155 Masson's Trichrome Stain, for connective tissue . . 156 Methyl Blue - Eosin (Mann) for demonstrating the vari- ous types of the cells in the anterior lobe of the pituitary 157 Methyl Green for amyloid .. .. .. .. ..158 Methyl Green - Pyronin (Pappenheim-Unna) for plasma ceils .. •• •• •• ••. •• •• 5 Methyl Violet 6B for amyloid . . . . . . . . 159 Methyl Violet - Metanil Yellow, for typhus fever rickettsiae in lungs of mice . . . . . . . . 160 Methyl Violet - Pyronin - Orange G (Bonney's Triple Stain) for chromatin, connective tissue, keratin, etc* •• •• •• •• •• •• ..1 uu Methylene Blue - Basic Fuchsin, rapid method of dem- onstrating Negri bodies .. ^.. .. .. .. 161 Methylene Blue - Basic Fuchsin for rickettsia . . . . 162 Methylene Blue Polychrome (Unna) for mast cells . . 163 Methylene Blue Polychrome - Glycerine Ether (Unna) for differentiating mast cells and plasma cells . . . . 164 MucicARMiNE - Metanil Yellow - Haematoxylin for mucin and connective tissue . . . . . . . . 165 Mucicarmine (Mayer) for mucin 166 XV CONTENTS Page 't>' MuciCARMiNE (Southgate) for mucin . . . . . . i66 MuciHAEMATEiN for mucus . . . . . . . . . . i66 Nadi Reaction for oxidase granules . . . . . . . . 1 67 Naphthol Blue Black - Haematoxylin - Brilliant Pur- PURiN - AzoFUCHSiN, for collagen, smooth muscle, etc. 168 Naphthol Green B - Haematoxylin (Weigert) for con- nective tissue . . . . . . . . . . . . 169 Neutral Red - Fast Green for staining both Gram-positive and Gram-negative organisms in sections . . . . 170 Nile Blue Sulphate for demonstrating fatty acids and neutral fats . . . . . . . . . . . . 171 Nile Blue - Picro Fuchsin (Murray-Drew) for bacteria and actinomyces in pathological tissues . . . . 172 Orange G - Crystal Violet (Bensley) for secretion ante- cedents of serous or zymogenic cells . . . . . . 173 Orcein for elastic fibres and connective tissue . . . . 174 Orcein - Aniline Blue - Orange G, a differential stain for elastic fibres, collagen and keratin . . . . . . 175 Orcein - Aniline Safranin for elastic and connective tissue fibres . . . . . . . . . . . . 176 Orcein - Giemsa Stain for syphilitic tissue . . . . 177 Orcein - Picro Fuchsin (Van Gieson) for elastic and colla- gen fibres . . . . . . . . . . . . . . 179 OsMic Acid, a rapid technique for staining fat in frozen sections . . . . . . . . . . . . . . 179 Pasini's Stain (Improved) for differentiation of connective tissues . . . . . . . . . . . . . . 180 Periodic Acid - Celestin Blue, for human and animal pituitary glands, demonstrating both muco-protein precuspors of the gonadtrophins ., .. .. 181 Periodic Acid - Feulgen Fuchsin (Hotchkiss) for Poly- saccharide structures . . . . . . . . • • 183 Phloxin - Haematoxylin for Hyaline in animal tissues 185 Phloxin - Methylene BIue, rapid smear technique for Negri bodies in brain tissue (j. R. Dawson's method) . . 186 Phloxin - Methylene Blue - Azur B, a very rapid modification of Mallory's original technique for normal and pathological tissues . . . . . . 187 xvi CONTENTS Page Phloxin - Tartrazine {A. C. LendrunCs technique)^ a gen- eral histological stain and for the demonstration of inclusion bodies . . . . . . . . . . . . i88 PiCRO - NiGROSiN for Eleidin and Keratin . . . . . . 189 Protargol - Gallocyanin (Foley) for nerve fibres, sheaths and cells . . . . . . . . . . . . . . 190 PuRPURiN for calcium deposits in pathological tissues . . 192 Quincke Reaction for Haemosiderin . . . . . . 193 Rhodamine B - Aniline Methylene Blue for splenic and lymphoid tissues . . . . . . . . • • i93 Saffron for connective tissue. . . . . . . . . . 194 Safranin - Crystal Violet - Fast Green - Orange 2 (S. S. Kalter's Quadruple Stain) . . . . . . 195 Safranin - Water Blue (Unna) for Collagen fibres . . 197 Scarlet R - Ethylene Glycol, an improved technique for staining fat . . . . . . . . . . . . 1 97 Scarlet R for staining fat . . . . . . , . . . 198 Silver Carbonate - Aniline Blue - Fast Green, for reticulin, elastin and collagen . . . . . . . . 199 Silver Nitrate - Gold Chloride - Paracarmine (Da Fano) for Golgi apparatus . . . . . . . . 202 Silver Nitrate - Hydroquinone for the detection of gold in fixed tissues of experimental animals . . . . 203 Sudan Black (J. R. Baker's technique) for lipids . . . . 204 Sudan Black, a specific stain for neutral fats . . . . 207 Sudan Black - Ethylene Glycol, an improved technique for lipid staining . . . . . . . . . . . . 208 Sudan Blue for demonstrating degenerated myelin . . 209 Sudan Brown for acute fatty degeneration not shown by Scarlet R . . . . . . . . . . . . . . 209 Sudan 2 for degenerating and intact myelin and fat . . . . 210 Thionin (Ehrlich) for mucin 211 Thionin (Ehrlich) for the differential staining of entamoeba 212 Thionin (Ehrlich) for nerve cells . . . . . . . . 213 Thionin for demonstrating malignant cells in biopsy material 213 ToLuiDiNE Blue for mucus . . . . . . . . . . 214 Trichrome Stain (G. Gomori) for connective tissue, etc. 214 Trichrome Stain (Masson), modified for pituitary gland, epithelium, thyroid nerve (normal and .tumour) etc. . . 215 B xvii CONTENTS Page Urea Silver Nitrate, for nerve fibres and nerve endings 217 Verhoeff's Stain for elastic fibres, nuclei and collagen . . 218 Water Blue - Orcein - Safranin for demonstrating epith- elial fibres . . . . . . . . . . . . . . 220 Weigert - French Elastin Stain (Moore's modification) for elastic tissues . . . . . . . . . . . . 221 Weigert - Pal Technique for myelin in brain and spinal cord and for peripheral nerves and ganglia . . . . 223 Wool Green - Haematoxylin - Ponceau S. for connec- tive tissue and muscle . . . . . . . . . . 224 Wright's Stain for general differentiation of blood cor- puscles ; for malarial parasites ; trypanosomes, etc. . . 225 SECTION 3— BOTANICAL METHODS Normal and Infected Tissues {a) GENERAL TREATMENT OF TISSUES . . . . 229 (h) MISCELLANEOUS MICROCHEMICAL TESTS 230 Aldehydes — Aleurone — Amygdalin — Amylodextrine — Anthocyanin — Arbutin — Asparagine — Calcium — Calcium Oxalate — Callose — Carotin — Cellulose — Chitin — Chlorides — Chlorophyll — Formic Acid — Glutathione — Inulin — Iodine — Iron — Lecithin — Nitrates — Pectic Substances — Phosphates — Phyto- sterol — Potassium — Proteins — Saponin — Sodium — Sulphates — Tyrosin. {c) STAINING TECHNIQUES 247 Acid Fuchsin - Aurantia for differentiating between bac- teria and mitochondria in infected tissues . . . . 247 Acid Rubin - Aurantia - Toluidine Blue (KuU's Stain), for starch grains and mitochondria . . . . . . 248 Aniline Hydrochloride, a rapid method for demonstrating lignified tissues . . . . . . . . . . . . 249 Basic Fuchsin, Ammoniacal for lignified walls and cutin . . 249 Borax Carmine (Grenacher) for bulk staining and for small whole mounts . . . . . . . . . . . . 250 xviii CONTENTS Page Chlorazol Azurine, a non-fading simple double stain suit- able for class work . . . . . . . . ..251 Chlorazol Black, a general-purpose stain . . . . 252 Chlorazol Paper Brown, B, a general-purpose stain . . 253 Cotton Blue - Safranin, for fungal hyphae in woody irXOoLX^o •• •• •• •• •• •• •• ^ 1^" Cyanin - Erythrosin, for cellulose and lignified tissues . . 255 Delafield Haematoxylin - Cellosolve, a general-pur- pose stain . . . . . . . . . . . . . . 255 Erythrosin - Lactophenol, a general-purpose stain . . 256 Gram's Iodine, a general-purpose stain . . . . 256 Haematoxylin (Heidenhain) - Aniline Blue (J. G. Vaughan's technique), for differential staining of nuclei, cytoplasm and cell walls of angiosperm shoot apices . . 257 Haematoxylin - Bismark Brown, for phloem tissues of woody plants . . . . . . . . . . . . 259 Heidenhain Haematoxylin - Safranin, a general-purpose stain useful for the demonstration of histological and cytological structures, as well as for algae, fungi, etc. . . 260 Iodine Green - Acid Fuchsin, for lignified tissues and for chromosomes . . . . . . . . . . . . 261 Johansen's Quadruple Stain, a general stain, excep- tionally good for sections of lichens . . . . . . 262 Johansen's Quintuple Stain, a general-purpose stain, particularly useful for leaves, roots, stems and ovaries 264 Lacmoid - Tannic Acid - Ferric Chloride, for phloem and contigous tissues . . . . . . . . . . . . 266 Lacmoid - Martius Yellow for callose in pollen tubes . . 268 LiGNiN Pink, a non-fading stain, specific for lignin . . . . 269 Magdala Red - Fast Green, for the differential staining of host tissues and parasites . . . . . . . . 269 Methyl Green - Phloxin - Glycerin Jelly, for the simultaneous double staining and mounting of pollen grains, differentiating functional from abortive grains 270 Periodic Acid - Feulgen Fuchsin (Hotchkiss) for poly- saccharide structures . . . . . . . . . . 183 Phloroglucinol for lignin, particularly suitable for hydro- phytes . . . . . . . . . . . . . . 271 xix CONTENTS Page Polyvinyl Lactophenol, for embedding brittle specimens of wood for sectioning . . . . . . . . . . 272 Safranin - Acid Fuchsin - for spermatozoids, zoopores etc. 276 Safranin - Aniline Blue, for gymnosperm ovules, arche- gonia, embryos, angiosperm stems and roots, etc. . . 273 Safranin - Aniline Blue for cellulose walls, cytoplasm, and for chromosomes . . . . . . . . . . 274 Safranin - Dianil Blue G, for differential staining of peronosporaceae . . . . . . . . . . . . 274 Safranin - Fast Green, FCF, a non-fading, general-pur- pose stain . . . . . . . . . . . . . . 277 Safranin - Fast Green, FCF - Cellosolve, a rapid, non-fad- ing stain in place of Sanfranin Light Green - Clove Oil 275 Safranin - Light Green - Cellosolve, a rapid general- purpose stain . . . . . . . . . . . . 278 Safranin - Light Green - Clove Oil, a general-purpose srain .. .. .. .. .. .. ,. 270 Safranin - Picro Aniline Blue, a rapid and simple method of demonstrating hyphae in wood sections . . 279 Safranin - Tannic Acid - Fast Green, FCF for roots and stems . . . . . . . . . . . . . . 280 Scarlet R or Sudan 3 - Ethylene Glycol, an improved technique for staining fat . . . . . . . . 281 Scarlet R for demonstrating fats . . . . . . . . 282 ScHULZE Solution (Chlor Zinc Iodine) for cell walls, proteins and starch . . . . . . . . . . 282 Tetrazolium Salt, for testing the viability of seeds . . 283 Thionin - Orange G for infected plant tissues, for chromo- somes, etc. . . . . . . . . . . . . 283 Titan Yellow^ for detection of magnesium in cells . . 284 Trypan Blue, a nuclear stain for plant material . . . . 285 SECTION 4— CYTOLOGICAL METHODS Acid Fuchsin - Toluidine Blue - Aurantia for mito- chondria . . . . . . . . . . . . . . 289 Acid Fuchsin - Picroindigo Carmine, for plant tissues as a cytological stain for root tips. . . . . . . . . 290 Alizarin Red, S (Benda's method) for mitochondria, etc. 291 Aniline Fuchsin - Iodine Green for mitochondria . . 292 XX CONTENTS Page Aniline Fuchsin - Picric Acid (Altmann) for mitochondria 293 Aniline Safranin (Babe's) for mitosis in animal cells . . 294 Basic Fuchsin - Picro Indigocarmine (Alcoholic) for plant tissues as a cytological stain for root tips . . . . 295 Basic Fuchsin - Picro Indigocarmine (Aqueous) for chromosomes in plant cells . . . . . . . . 295 Breinl's Triple Stain, for chromosomes . . . . . . 296 Copper Chrome Haematoxylin (Bensley), for mito- chondria . . . . . . . . . . . . . . 297 Cotton Red - Methyl Violet - Orange G for plant cells 299 Feulgen Stain for demonstrating mitosis in plant cells . . 300 Gentian Violet Picric Acid - Iodine for chromosomes in plant cells . . . . . . . . . . . . 300 Haematoxylin (Regaud) for mitochondria 302 Haematoxylin - Safranin, a cytological stain for plant cells 303 Iron Aceto Carmine (Belling) for chromosomes in micro- sporocytes . . . . . . . . . . . . 304 Methyl Green - Acid Fuchsin - Erythrosin, a cytological stain for plant cells . . . . . . . . . . 305 Methyl Green - Acid Fuchsin for chromosomes in plant WC^lld •• •• •• •• •• •• •• ^ Methyl Violet - Eosin Scarlet, a botanical stain for mitosis in root tips . . . . . . . . . . 307 NiGROSiN, for salivary chromosomes of Drosphila . . . . 307 Safranin - Crystal Violet (Hermann), for chromosomes 309 Safranin - Gentian Violet - Orange G {Flemming Tri- colour Stain) for chromosomes, etc. . . . . . . 309 Wright's Stain, for demonstrating cytoplasmic changes in plant cells . . .. .. .. .. .. ..310 SECTION 5— FLUORESCENCE MICROSCOPY {a) GENERAL INFORMATION 315 {b) EQUIPMENT REQUIRED 316 {c) STAINING METHODS 317 1. Cytoplasm, Nuclei, Nucleoli and Chromatin, Dif- ferential Staining of 318 2. Fat Staining .. .. 318 xxi CONTENTS Page 3. Intravital Staining . . 4. Muscle 5. Nerve Tissues, Differential Staining of 6. Soil Bacteria, Staining and Counting ; Differentia- tion OF Living from Dead Cells 7. Trypanosomes, Living, in Blood 8. Trypanosomes, in Dried Blood Films 9. Tubercle Bacilli 10. Virus Staining . . SECTION 6— HISTOCHEMICAL METHODS 319 320 321 322 323 324 325 326 Micro- Incineration for the detection and location of mineral salts .. .. .. .. .. ..331 3 -Hydroxy - 2-Naphthoic Acid - Tetrazotized o-Ani- sidine, for demonstrating sites of carbonyl activity . . 333 Metanil Yellow - Iron Haematoxylin for radioauto- graphs . . . . . . . . . . . . . . 334 Methyl Green - Pyronin - Ribonuclease (Brachet), for detecting ribonucleic acid and desoxyribonucleic acid in the same cell . . . . . . . . . . . . 336 Nile Blue Sulphate - Safranin, for phospholipids . . 340 Phosphomolybdic Acid - Eosin, for localization of choline containing lipids . . 341 Sudan Black 342 SECTION 7— SMEAR PREPARATIONS Albert's Stain (Layboum's modification) for diphtheria bacilli . . . . . . . . . . . . . . 345 Alcian Blue, for bacterial capsules and polysaccharides . . 346 Aniline Blue - Eosin B, for spermatozoa 347 Aniline Gentian Violet, a rapid stain for Treponema pallida . . . . . . . . . . . . . . 348 AzuR L for the detection and staining of epidermophytic infection . . . . . . . . . . . . . . 349 xxii CONTENTS Page Basic Fuchsin for Treponema pallida . . . . . . 349 Breed^s Stain for staining and counting bacteria in milk . . 350 Brilliant Cresyl Blue for reticulated cells, platelets, etc. 351 Carbol Crystal Violet for fibrin network in blood smears 352 Carbol Fuchsin for Treponema pallida and other spiro- cnaeres •• •• •• •• •• •• ' * jjo Carbol Fuchsin for tubercle bacilli and other acid-fast bacteria . . . . . . . . . . . . . . 354 Carbol Fuchsin - Borrel Blue for leprosy and tubercle bacilli . . . . . . . . . . . . . . 355 Carbol Fuchsin - Brilliant Yellow for tubercle bacilli in sputum . . . . . . . . . . . . . . 356 Casteneda's Stain, for rickettsiae and elementary bodies 356 Crystal Violet for bacterial capsules . . . . . . 357 Crystal Violet, a simple stain for spirochaetes . . . . 358 Dahlia, for Heinz bodies in blood smears . . . . . . 358 Dorner's Stain for spores . . . . . . . . • • 359 Ehrlich's Triacid Stain for blood 359 EosiN - Gentian Violet for basal bodies . . . . 360 EosiN - Methylene Blue - Basic Fuchsin, a general stain for bacteria, blood and spirochaetes . . . . . . 361 Field's Stain for thick blood smears for malarial parasites 362 Flagella Stain (Cesares-Gil) for colon and typhoid organisms . . . . . . . . . . . . . . 363 Fontana Stain for spirochaetae . . . . . . • • 3^3 Gentian Violet (Noland) a combined fixative and stain for protozoa . . . . . . . . . . . . . . 364 GiEMSA Stain for blood, malarial parasites, Treponema pallida^ trypanosomes, etc. . . . . . . • • 3^5 GiEMSA - May - Grunwald Stain for blood, malarial parasites, trypanosomes, etc. . . . . . . . . 368 Gram's Iodine - Aniline Gentian Violet, for spirochaetae of Vincent's Agina and for Trep. pallidum . . . . 371 Gram's Stain for bacteria 368 Gram's Stain (Jensen) for bacteria . . . . . . 369 Gram's Stain (Kopeloff and Beerman) for bacteria . . 370 Haematoxylin (Phosphotungstic, Mallory) for entamoeba 372 Haematoxylin (Weigert) - Bordeaux Red for permanent preparations of anophcline midgut . . . . . . 372 xxiii CONTENTS Page Haematoxylin - EosiN - Indigo Carmine for the differ- ential staining of vaginal smears . . . . . . 373 Iodine - Eosin for intestinal amoebae and flagellates . . 374 Janus Green, B for staining oocysts of coccidia in faeces . . 375 Janus Green - Neutral Red, for supravital staining of blood 376 Jenner Stain for the cytological examination of blood . . 377 Leishman Stain for blood, malarial parasites, trypanosomes, wLw* •• •• •• •• •• •• •• 1// Lugol's Iodine - Eosin for differentiating between spores and vegetative forms in bacterial smears . . . . 379 Macchiavello's Stain, for rickettsiae Malachite Green a simple method of staining spores . . 380 Malachite Green - Basic Fuchsin, for yeasts . . . . 381 Malachite Green - Pyronin - Crystal Violet (Sandi- ford), a contrast stain for gonococci and meningococci. . 381 May - Grunwald Stain for the cytological examination of blood . . . . . . . . . . . . . . 382 Methyl Green - Pyronin, for gonococci . . . . . . 382 Methyl Violet ioB, for the direct staining of elementary bodies . . . . . . . . . . . . . . 383 Methylene Blue - Carbol Fuchsin for flagella and cap- sules in bacterial films . . . . . . . . . . 384 Methylene Blue - Gentian Violet for gonococci . . 385 Methylene Blue - Safranin for polar bodies in bacteria 385 New Fuchsin - Congo Red, for bacterial cell walls, particu- larly for B. colt and B. cereus . . . . . . . . 386 Newman's Stain, a single solution for defatting and staining milk smears for counting bacteria . . . . . . 387 Nile Blue Sulphate for protozoa, yeasts, etc. . . . . 388 Papanicolaou Stain, a differential stain for vaginal smears 388 Peroxidase Stain (Cowdrey's method) for blood . . . . 389 PiCRO - Methyl Blue - Eosin for urinary casts . . . . 390 PiNACYANOL - Neutral Red, for Supravital staining of blood 392 Ponder's Stain (Kinyoun's modification) for the differ- entiation of metachromatic granules of diphtheria organisms . . . . . . . . . . . . . . 393 Pyronin - Alphanaphthol (Graham) for oxidase granules in blood . . . . . . . . . . . . . . 394 XXIV CONTENTS Safranin - Light Green for spirochaetes, etc. . . Schorr's Stain for vaginal smears . . Schorr's Stain (single solution) for vaginal smears Sudan Black - Eosin - Methylene Blue for lipoid granules in leucocytes Sudan Black - Safranin for demonstrating fat in bacteria Sudan 3 for blood in the spinal fluid, differentiating fresh haemorrhage from old Sudan 3 for staining fat in faeces Tetrachrome Stain (MacNeal) for differentiating types of leucocytes . . Thionin (Ehrlich), a rapid stain for rickettsia in smears . . Vaginal Smear Stain, M.F.4 for the rapid staining of vaginal smears, sharply contrasting cornified from non- cornified cells Vaginal Smear Stain, PX, a differential stain for vaginal smears Victoria Blue 4R for elementary bodies . . Victoria Blue 4R for Treponema pallida . . Victoria Blue - Kernechtrot Red - Light Green, for elementary bodies Wright's Stain for general differentiation of blood cor- puscles ; for malarial parasites, etc. Wright's Stain for demonstrating Trichomonas riedmuller APPENDIX Atomic Weights, Tables of Conversion Tables Formulae (Fixatives) Formulae (Stains and Reagents) Refractive Indices Saturated Solutions (Reagents) 7. Specific Gravities 8. Stain Solubilities and Molecular Weights I. 2. 3- 4- 5- 6. Page 395 395 396 397 397 398 399 400 400 401 402 402 403 404 405 406 407 409 410 411 412 426 428 430 430 437 XXV SECTION I— GENERAL METHODS FIXATION The main objects of fixation are: (a) To kill the cells suddenly and uniformly so that they retain, as near as possible, the same appearance which they possessed in life. (b) To preserve the tissues, cells, etc., by the inhibition of putrifactive and autolytic changes. (c) To set and hold intra-cellular bodies, cells, etc., by precipi- tation in the positions which they occupied in life, thereby facili- tating the closest possible study of the histology and cytology of the cells. (d) To facilitate differentiation in the refractive indices of cer- tain cell elements which would otherwise be invisible owing to the exceedingly narrow margin betw^een the refractive index of one type of cell and that of other types. (e) To render the cells and tissue constituents resistant to the subsequent processes such as dehydration, clearing, embedding, staining, etc., prior to their examination under the microscope. (/) To facilitate proper staining of tissues. Here it should be mentioned that some fixatives act as mordants while others act as inhibitors for certain stains, and it is, therefore, of considerable importance that a suitable fixative should be employed for a particular staining technique or a particular staining technique should be chosen to suit material which has already been treated with a particular fixative. Recommendations as to suitable fixa- tives are given for most, if not all, of the staining techniques des- cribed in this book. It is essential, if good results are to be achieved, that tissues should be removed from the body with the least possible delay and fixed immediately. Bacteria, protozoa and other unicellular organisms should be living at the instant of fixation. The follow- ing points must be observed in order to avoid failure, waste of time, effort and materials : MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 1 . Pieces of tissue should, whenever possible, be cut into slices 2 to 6 mm. in thickness to permit penetration of the fixative throughout within a reasonable time. 2. The container in which the material is to be fixed should be of sufficient size to take the pieces of tissue without their folding or bending. 3. If large organs are to be fixed, large incisions should be made to allow thorough penetration of the fixing fluid. 4. A volume of the fixing fluid roughly about twenty to fifty times as great as that of the material to be fixed is necessary. 5. Material must not be left in the fixing fluid beyond the necessary time, but should be taken out, washed, dehydrated, cleared and embedded or stored in a fluid, suitable for the parti- cular material, until it is required for embedding. 6. After fixation, and before proceeding to dehydration, careful washing out of the excess fixative is necessary, except in the case of alcohol which requires no washing out. In most cases running water is employed for this purpose, but for some tissue cells and cell constituents, and for some fixatives, alcohol must be used. In all cases, however, it is necessary to use liberal quantities of the liquids for washing out. 7. A fixative suitable to the material to be examined and com- patible with the stains to be employed should be chosen, as dis- regard of this factor will, as previously stated, lead to failure and disappointment as well as waste of time, effort and materials : the importance of this rule cannot be over-emphasized. (a) FIXATIVES The number of these from which to choose is legion, although the number in everyday use is comparatively small. Details of some of the more commonly used of these are given below : for- mulae of others are given in the appendix. Acetic Acid, Glacial Recommended for : Rapid fixation of strongly contracting organisms. 4 SECTION ONE Technique: Fix in the warm acid for a maximum period of fifteen minutes; then remove excess acid by washing in 30-50% alcohol. Remarks: Acetic acid glacial is rarely used alone : it causes the swelling of cell constituents, etc., and it is of most value when mixed with such substances as formalin, alcohol, mercuric chloride, etc., to counter- act their shrinking effect. Acetone Recommended for : Rapid fixation of brain tissue for rabies diagnosis. (R. D. Lillie, Histopathologic Technique.) It is also employed for fixing tissue enzymes, particularly phosphatases and lipases. Technique: Thin slices of tissue are fixed in pure acetone for twelve to twenty-four hours at 0° C. ; they are then dehydrated by immer- sion in two changes of pure acetone for two hours in each at room temperature, and afterwards cleared by immersing for half an hour in each of two changes of benzene before infiltration with paraffin wax.* Alcohol Absolute Recommended for : Glycogen, Amyloid, Fibrin, Hyaline, Haeipofuscin, Phos- phatase. Technique: Fix from two hours to several days according to the nature of the material; dehydrate; clear. Remarks: Unsuitable for fats and lipines as these are dissolved by the higher concentrations of alcohol. *Gomori, Proc. Soc. Exp. Biol, and Med., 58, 362, 1945. 5 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Allen's Fixative (B.15) Recommended for : Chromosomes; plant tissues, particularly buds. Formula: Picric acid, saturated, aqueous . 75 ml. Formalin (formaldehyde 40%) . . 20 ml. Acetic acid, glacial . . 5 ml. Chromic acid . . . 1-5 gm ^^ X W^ •• •• •• •• . . 2 gm. Technique: Fix tissues from four to sixteen hours then immerse in frequent changes of 70% alcohol over a period of forty-eight hours or until the yellow coloration due to picric acid ceases to come out. Remarks: The solution deteriorates very rapidly and it is, therefore, essen- tial that this fixative should be prepared only when it is required for immediate use. Bouin's Fluid Recommended for : Embryological specimens, for elementary bodies, Purkinje cells, Argentaffin reaction, and for animal tissues in general. Formula: Picric acid, saturated aqueous . . 75 ml. Formalin . . . . . . . . 25 ml. Glacial acetic acid . . . . • • 5 ml. Technique: Fix from eighteen hours to two days; then wash in 50% fol- lowed by 70% alcohol until most of the yellow coloration, due to picric acid, is extracted. Alternatively the picric acid can be washed out with the alcohols after the tissues have been embedded and sectioned. It is not essential that all the picric acid, which serves as a 6 SECTION ONE mordant enhancing many staining effects, should be entirely extracted from the fixed tissues. Remarks: This fixative, which keeps indefinitely and causes only slight shrinkage of tissues, is compatible with almost every staining technique : it is considered to be a valuable fixative for most pur- poses, although it is unsuitable for kidney and mucin. Its pene- tration power is great, and delicate material should be left in contact with this fixative only for the minimum time, to avoid over-fixation : this applies to cytological work in particular. There are many modifications of Bouin, of which Allen's Fixative, B.15, has proved to be the most satisfactory for chromosomes in mammalian tissues. Bouin - Duboscq (Duboscq-Brasil, or Alcoholic Bouin) Recommended for : Arthropods containing parasites and protozoan cysts, and for chitinous tissue. Formula: Absolute alcohol . 48 ml. Formalin (formaldehyde 40%) . 30 ml. Glacial acetic acid . 7*5 ml. Picric acid, saturated aqueous . 12-5 ml Distilled water . 15 ml. Technique: Fix from eighteen hours to two days; then wash in 70% alcohol. Remarks: This is stated to be more penetrating than aqueous Bouin and for this reason it is employed for hard tissues. Carnoy's Fluid Recommended for : Glycogen ; for animal tissues in general, and for plant cytology. c 7 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Formula: Absolute alcohol . . . . . . 60 ml. Chloroform . . . . . . . . 30 ml. Glacial acetic acid . . . . . . 10 ml. Technique: Animal Tissues. Fix for one and a half to two hours; then transfer to two changes of absolute alcohol before clearing and embedding. Plant Tissues. Fix root tips for a quarter-of-an-hour ; anthers, for one hour. Remarks: A rapid fixative with great penetrating power. Carnoy - LeBrun Fluid Recommended for : Insects and ticks, and as a rapid and penetrating fixative for plant tissues. Formula: Absolute alcohol . . . . . . 30 ml. Glacial acetic acid . . . . • • 30 ml. Chloroform . . . . . . . . 30 ml. Mercuric chloride . . to saturation (about 10 gm.) Technique: Fix from half to one minute ; then wash in 95% alcohol. Remarks: Not reconmiended for routine fixation of plant tissues. Champy's Fluid Recommended for : Plant and animal tissues in general; mitochondria and other cytological detail. Formula: Potass, dichromate 3% aqueous . . 35 ml. Chromic acid 1% aqueous . . . . 35 ml. Osmic acid 2% . . . . . . 20 ml. 8 SECTION ONE Technique: Fix for six to twenty-four hours; then wash in running water for the same length of time. Flemming's Fluid (Strong) Recommended for : Plant and animal tissues, for cytological detail; for cellular structures; and for fat. Formula: A. Chromic acid I % .. .. 30 ml. Acetic acid, glacial . . . . 2 ml. B. Osmic acid 2% . . . . . . 8 ml. Mix A and B immediately before use. Technique: Fix for one to twenty-four hours according to the material ; for chromosomes an hour is sufficient. Wash with running water for twenty-four hours. Remarks: The penetration power of this fixative is poor ; and that of the weak solution {see below) is poorer still; however, either solution gives good results with basic stains, particularly safranin, and iron haematoxylin. Quite apart from their high cost, Flemming fixa- tives should definitely not be employed as general fixatives, but only in special cases for very small pieces of tissue where fixation extending through a layer of about four or five cells in thickness is sufficient, as this is the limit of their penetrating power even in loose-celled tissues. Flemming's Solution (weak) Recommended for : All purposes for which Flemming's strong solution is used. Formula: A. Chromic acid 1% . . . • 25 ml. Acetic acid 1% . . . . . . 10 ml. B. Osmic acid 1% . . . . . . 10 ml. Distilled water . . . . • . 50 ml. MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: Used in the same way as the strong solution {see above), except that the volume of the fixative required is about eight or ten times that of the material to be fixed. Remarks: See under Flemming's Fluid (strong). Formalin Neutral Recommended for : Animal tissues in general, particularly nervous tissue. Formula: Formalin (formaldehyde 40%) . . 100 ml. Tap water . . . . . . . . 900 ml. Magnesium carbonate . . . . to excess Shake well and allow to stand several hours at least; then decant off the volume of the clear fluid required for fixation. Technique: Fix at least twenty-four hours at room temperature, or six to eight hours at 50-60° C. Washing out is unnecessary. Remarks: Formalin penetrates well, tissues may be kept in it for long periods without undue hardening, although there is a gradual decrease in basophilia of cytoplasm and nuclei, and certain cyto- plasmic structures are not hardened by it sufficiently to permit paraffin embedding. Best adapted to material which is to be em- bedded in celloidin rather than in paraffin wax; also suitable for frozen sections. Formalin Buffered Formula: Neutral formalin, as above . . . . i litre Sodium dihydrogen phosphate, mono- hydrate, A.R. . . . . • • 4 gni. Disodium phosphate anhydrous, A.R. 6-5 gm. Technique: As for neutral formalin. 10 SECTION ONE Remarks: Neutral formalin turns acid on keeping owing to the production of formic acid, whereas the buffered fixative remains neutral. Recommended for : All purposes for which neutral formalin is employed where a neutral fixative is required. Formol - Alcohol Recommended for : Glycogen in animal tissues. Fibrin. Peroxidase. Plant tissues, particularly pollen tubes in styles. Formula: Formalin (40% formaldehyde) . . 100 ml. Alcohol 70% 900 ml. Technique: Fix for three to six hours; then dehydrate, clear and embed. Alternatively, if it is not convenient to dehydrate, clear and embed at once, the tissues may be stored for long periods without deleterious effects in 70% alcohol. Remarks: This fixative, which penetrates quickly, while compatible with most stains, is particularly suitable for indigo carmine. Kelly's Fluid Recommended for : Animal tissues in general, but particularly for blood-forming organs. Formula: Potassium dichromate ... . . 5 gm. Mercuric chloride Sodium sulphate crystals Formalin (40% formaldehyde) Distilled water 10 gm. 2gm. 10 ml. 200 ml. N.B. — The formalin should not be added until the fixative is required for immediate use. II MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: Fix for twelve to twenty-four hours. Wash in running water for the same time. Transfer to 80% alcohol; dehydrate; clear and embed. Remarks: Not recommended for bacteria, cytoplasm degeneration, necrosis or regeneration. Hermann's Fluid Recommended for : Cytological work. Formula: Platinic chloride 10% . . . . 6 ml. Osmic acid 1% . . . . • • 32 ml. Glacial acetic acid . . . . • • 4 nil. Distilled water . . . . . . 38 ml. N.B. — The solution should be freshly prepared. Technique: Fix from twelve to sixteen hours, wash in running water for three to six hours ; then treat as for Flemming fixed-material. Remarks: While this fixative mordants chromatin for staining with basic stains, it inhibits staining with acid stains. Good plasma staining is difficult if not imposssible after this fixative. Rleinenberg's Fluid Recommended for : Embryos; marine organisms, arthropods, chitinous material. Formula: Sulphuric acid 1% . . . . . . 100 ml. Picric acid, saturated aqueous . . 49 ml. 12 SECTION ONE Technique: Wash out the picric acid with warm 70% alcohol, followed by increasing strengths of alcohol. Remarks: This fixative is a powerful penetrant of chitin. Lewitsky's Fluid Recommended for : Plant cytology. Formula: Formalin (formaldehyde 40%) . . 100 ml. Distilled water . . . . . . 100 ml. Chromic acid 5% aqueous . . . . 100 ml. Technique: Fix tissue for twelve to twenty-four hours; then wash for six to sixteen hours in running water. Marchi's Fluid Recommended for : Animal and plant tissues generally. Formula: Potassium dichromate 2-5% aqueous 100 ml. Sodium sulphate crystals . . . . i gm. Osmic acid 1% aqueous . . • • 50 ml. Note. — The last item should be added immediately before use. Technique: Immerse thin pieces of tissues, not more than 2 mm. thick in the fixative for four to eight days ; then wash in running water for twelve to sixteen hours. Transfer to 70% alcohol, afterwards dehydrating, clearing and embedding in the usual way. Remarks: This fixative is also employed to blacken nerve fibres. Navashin's Fluid Recommended for : Cytological study of plant tissues. 13 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Formula: Chromic acid . . .. .. .. 1-5 gm. Acetic acid 10% aqueous . . . . 100 ml. Distilled water . . . . . . 60 ml. Formalin (formaldehyde 40%) . . 40 ml. Note. — The formaldehyde should not be added until the solu- tion is required for immediate use. Technique: Fix material for one to two days; then wash twelve to sixteen hours in running water. Dehydrate ; clear and embed. Orth's Fluid Recommended for : Demonstration of acute degenerative processes to be stained with Giemsa, Wright, or Leishman Stain, and for Intestine. Formula: Potassium dichromate . . . . 2-5 gm. Sodium sulphate crystals . . . . i gm. Distilled water . . . . . . 100 ml. Formalin (formaldehyde 40%) . . 10 ml. The last item should not be added until the fixative is required for immediate use. Technique: Fix pieces of tissue up to i cm. in thickness for two to four days ; then wash in running water from twelve to twenty-four hours. Transfer to 80% alcohol; dehydrate; clear and embed. Remarks: This fixative may be employed as a general fixative for animal tissues. It is useful where a firm consistency of tissue is required, but it is not recommended for sharp histological detail. The value of the sodium sulphate in the above formula is extremely doubtful and it appears that this constituent may be left out without any noticeable effect. 14 SECTION ONE Petrunkevitch's Cupric Paranitrophenol Fixative* Recommended for : Tissues in general. Formula: Alcohol 60% 100 ml. Nitric acid, pure (sp. gr. i*4i — 1-42) 3 ml. Ether . . . . . . . . 6 ml. Cupric nitrate Cu(N03) 2, 3H2O . . 2 gm. Paranitrophenol pure, cryst. . . • • 5 gm. Technique: Fix material for twelve to twenty-four hours. Wash in 70% alcohol; dehydrate; clear and embed. Remarks: All stains conmionly in use may be employed after this fixative, which causes less hardening than most other fixatives. Regaud's Fluid Recommended for : Mitochondria and rickettsia in animal tissues. Formula: Potassium dichromate . . • • 3 gn^- Distilled water . . . . . . 100 ml. Formalin (formaldehyde 40%) . . 25 ml. Technique: Fix material for three days changing the fluid every day. Im- merse in 3% potassium dichromate for six to eight days; then wash in running water for twenty-four hours. Dehydrate; clear and embed. Remarks: Suitable for Giemsa stain and for Masson's trichrome stain. Schaudinn's Fluid Recommended for : Animal tissues in general. Protozoa. •A. Petrunkevitch, Science 77, 1 17-18, 1933. 15 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Formula: Mercuric chloride saturated aqueous 66 ml. Absolute alcohol . . . . . . 33 ml. Glacial acetic acid . . . . . . i ml. N.B. — The acid should be added immediately before use. Technique: Fix tissues for six to sixteen hours. Wash in several changes of 70% alcohol. Remarks: The fixative may be used at about 65° C. when less time is required for its penetration. Susa Fluid (Heidenhain) Recommended for : Animal tissues in general. Formula: Mercuric chloride, saturated aqueous 50 ml. Trichloracetic acid . . . . . . 2 gm. Formalin (formaldehyde 40%) . . 20 ml. Distilled water . . . . . . 30 ml. Glacial acetic acid . . . . • • 4 nil. Technique: Fix for five to twelve hours; then wash out with 95% alcohol. Remarks: Compatible with most stains, but not with Weigert's elastin stain. Susa offers the advantage over most other fixatives in that it causes less shrinkage and less hardening, thereby rendering tissues easier to cut. Zenker's Fluid Recommended for : Perfect histological detail in animal tissues in general. 16 SECTION ONE Formula: Potassium dichromate 2-5% aqueous 100 ml. Mercuric chloride . . . . • • 5 gin- Glacial acetic acid . . . . . . 5 ml. N.B. — The acid should not be added until the fixative is required for immediate use. Technique: Immerse slices of tissue in the fluid for six to twenty-four hours according to the nature of the material and the thickness of the slices. Wash in running water for twelve to twenty-four hours; then transfer to 80% alcohol. Remarks: Unsuitable for Mitochondria. Suitable for Mallory's connec- tive tissue stain; for demonstration of Muscle, Fibrin, Haemo- fuscin, Purkinje cells, etc. (b) DEHYDRATION, CLEARING, EMBEDDING, SECTIONING, ETC. CELLOIDIN METHOD FOR EMBEDDING TISSUES For preserving the relations of cell layers of different con- sistency, as are contained in the eye; for large objects; for pieces of central nervous system ; and for hard tissues such as decalcified bone. Solutions required: A. Celloidin 8% Celloidin flakes . . . . • • 25 gm. damped with absolute alcohol* Absolute alcohol . . . . , . 150 ml. Ether . . . . . . . . 163 ml. *Note: This may be obtained in 25-gm. bottles, ready damped with absolute alcohol. 17 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Pour the alcohol and ether into a clean, absolutely dry, wide-mouth bottle of about 32 ounce capacity, and mix by inverting the bottle several times, re- leasing the stopper at intervals, to release the pressure of ether vapour; then add the Celloidin flakes and invert the bottle as before. Leave for about 12 hours to dissolve ; inverting or shaking the bottle gently at intervals. B. Celloidin 4% Celloidin solution 8% . . . . 2 volumes Absolute alcohol . . . . . . i volume Ether i volume Mix as described above, in a large-stoppered wide- mouth bottle. C. Celloidin 2% Celloidin 4% Absolute alcohol . . Ether D. Cedarwood oil, for clearing Chloroform 2 volumes I volume I volume I volume I volume Technique: (a) Wet Method 1. Pieces of tissue not thicker than 5 mm. are fixed in the usual manner. 2. Wash in running water for the prescribed time for the parti- cular fixative. If a fixative containing mercury has been used, remove mercurial precipitate by the standard technique. 3. Immerse the pieces of tissue for two hours in each of the following: 50%, 70% and 90% alcohol. 4. Immerse in absolute alcohol from two to sixteen hours, according to the nature and thickness of the tissue. 5. Immerse for twenty-four hours in a mixture consisting of equal volumes of absolute alcohol and ether, in a stoppered wide- mouth bottle, which must be absolutely dry. 6. Impregnate with 2% Celloidin solution from five to seven days. 18 SECTION ONE 7. Transfer to 4% Celloidin for five to seven days. 8. Impregnate with 8% Celloidin for three or four days. 9. The tissue is then taken out of the Celloidin and put into a mould made by folding a piece of writing paper, and the whole is then placed in a desiccator and left for several days, lifting the desiccator lid for a few seconds each day to accelerate the harden- ing of the Celloidin. If, through shrinkage of the Celloidin during this process, the tissue becomes exposed, pour on more Celloidin solution to cover it. Hardening of the block may be hastened by placing 1-2 ml. of chloroform in the bottom of the desiccator. The block is hard enough for sectioning when no impression is left after pressing with the ball of the thumb. 10. The base of the hardened Celloidin block is dipped into 8% Celloidin then fixed to a roughened wooden or a vulcanite block by pressing firmly, afterwards leaving for at least half an hour with a weight on top. 11. Expose to chloroform vapour for half an hour; then attach the wooden or vulcanite block to the microtome holder, or store the Celloidin block mounted on the wooden or vulcanite block in 80% alcohol until required for sectioning. 12. The microtome knife and the Celloidin block must be kept moist with 70% alcohol and each section as it is cut must be trans- ferred by means of a camel-hair brush, moistened with 70% alcohol, into a suitable vessel containing 70% alcohol in which the sections can be stored indefinitely until required for staining. 13. When required for staining the sections should be removed from the 70% alcohol by means of a small camel-hair brush, or a piece of thin glass rod bent at one end, and transferred to a series of watch glasses containing the reagents and stains, arranged on the bench in the order in which they are to be used. For instance, if it is desired to stain the sections with Haematoxylin and Eosin, the steps are as follows : 14. Immerse sections in 50% alcohol for a few minutes; then transfer to water. 15. Stain with Ehrlich Haematoxylin by the standard technique. 16. Blue in tap water ; then stain in Eosin (aqueous solution). 19 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 17. Transfer to 70% alcohol; then immerse for five minutes in each of two lots of 96% alcohol. Note: Absolute alcohol must be avoided as Celloidin is dis- solved by it. 18. Immerse for five minutes each in two lots of Carbol-Xylol. 19. Pass into two changes of xylol. 20. Mount in balsam or Dammar-Xylol. {b) Dry Method 1. Proceed exactly as described above up to and including step No. 8 ; then take the tissue out of the Celloidin and put it into a paper mould as described in step No. 9 (above). 2. Place the block in a desiccator for a day, lifting the lid for a few seconds every hour or so; then leave in the desiccator over- night. 3. Next morning place the block in a mixture consisting of equal volumes of cedarwood oil and chloroform and add another 8 vol- umes of cedarwood oil, one volume at a time, every hour for small objects, or every day in the case of large objects. The Celloidin should now be wholly transparent. 4. Fix the Celloidin block to the wooden or vulcanite block as described in step 10 (above). 5. Sections may now be cut without the necessity of moistening the knife or the block. Note: Blocks prepared by this method are stored in a dry wide- mouth stoppered bottle. CELLOIDIN - PARAFFIN WAX (Double Embedding) For serial sections embedded in Celloidin Solutions required: Celloidin 1% in Methyl Benzoate This is prepared by adding the appropriate weight of air-dried Celloidin flakes to a quantity of Methyl Benzoate in a clean dry 20 SECTION ONE corked flask or bottle. Shake well; allow the flask or bottle to stand upright for an hour or so ; then leave it inverted for an hour, afterwards leave it lying horizontally for a time; then stand it upright again, and repeat the process at intervals throughout the day and leave the bottle lying on its side overnight : next morning, solution should be complete and it is only necessary to shake the bottle well to ensure a homogeneous solution. Technique: 1. Tissues are fixed and washed in running water, and any mer- curial precipitate removed in the usual manner. 2. Immerse for two hours in each of the following: 50%, 70% and 90% alcohol. 3. Transfer to absolute alcohol for two to sixteen hours. 4. Immerse in Methyl Benzoate - Celloidin solution for twenty- four hours, at the end of which time pour oflF the solution and replace with a fresh lot in which the tissue should remain for a further forty-eight hours : if the tissue is not now clear, transfer it to a fresh lot of Methyl Benzoate - Celloidin solution for a further period of seventy-two hours. 5. Immerse in three changes of pure benzene, for four hours in the first lot, eight hours in the second, and twelve in the third. 6. Transfer to a mixture consisting of equal parts of paraffin wax and pure benzene in the embedding oven for an hour. 7. Immerse in two changes of pure paraffin wax from a quarter of an hour to six hours in each, depending upon the thickness and the nature of the tissue. Note: It is of utmost importance that tissues should be kept in the embedding oven just long enough for the paraffin wax to penetrate fully. Prolonged heating in the oven causes shrinking and hardening of the tissues rendering sections difficult to cut. If, on the other hand, any of the Methyl Benzoate remains in the tissue and sufficient time has not been allowed for proper pene- tration of the paraffin wax satisfactory sections cannot be cut. It is best, whenever possible, to cut thin slices of tissue for embedding, preferably not more than 5 mm. thick so that the total time for impregnating in the pure paraffin wax need be no longer than three hours. Large objects such as whole embryos need a total 21 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES time of twelve hours in pure paraffin wax. Pieces of brain and spinal cord about 5 to 10 mm. thick, skin, and large objects such as whole embryos need at least three changes of pure paraffin wax for a total time of about 12 hours, whereas organs such as spleen, containing a large amount of blood, muscle, fibrous tissue, require no longer than a total of three hours in the paraffin baths. It is, however, only by experience that the technique of embedding can be mastered. 8. Cast the tissue in an embedding mould and proceed as in the case of paraffin sections {see page 32). CELLOIDIN - PYRIDIN A rapid method of dehydrating, clearing and embedding, which obviates the use of alcohols and the consequent hardening of tissues Reagents required: A. Pyridin, extra pure, redistilled . . i volume Celloidin 4% in equal volumes of absolute alcohol and ether . . i volume B. Celloidin 8% in equal volumes of absolute alcohol and ether. Technique: 1. Pieces of tissue are fixed in the usual manner. 2. Wash in running water for the prescribed time for the parti- cular fixative employed. If a fixative containing mercury has been used, remove mercurial precipitate by the standard technique. 3. Immerse the tissue in two changes of Pyridin, from two to eight hours in each, according to the nature and the thickness of the tissue. 4. Immerse for twenty-four hours each in two changes of a mixture consisting of equal volumes of Pyridin and 4% Celloidin (formula as above). 5. Immerse in 8% Celloidin for twelve hours. 6. The tissue is then removed from the Celloidin bath, blocked and cut into sections by the standard technique described on pages 19-20. 22 SECTION ONE FROZEN SECTIONS For the identification of fat in tissues ; for certain impreg- nation methods for the central nervous system; and for the rapid examination of pathological material, such as pieces of tumour, which may be sectioned, stained and diagnosed within a few minutes of their removal by the surgeon in the operating theatre Sections as thin as ^fj, may be cut, and an advantage of this method is that there is a lesser degree of shrinkage than in the case of paraffin-embedded material. It is not, however, possible to cut serial sections by this method, and sections cannot be stored before staining as in the case of paraffin-embedded material. Frozen sec- tions should be employed only for the specific purposes mentioned above and not as an alternative to paraffin embedding. It should be noted that frozen sections are manipulated in the same way as Celloidin sections, but greater care must be exercised on account of the absence of any embedding mass. Tissues should not be frozen too hard or the sections will curl up and split. A special microtome is required for cutting frozen sections. Rapid Method for Staining Fats Solutions required: A. Sodium chloride A.R. grade 0*9% in distilled water . . • • 95 n^l- Formalin . . . . . . • • 5 i^l- B. Sudan Black B, saturated in 70% alcohol (this should be freshly filtered). NOTE: Ethylene or Propylene Glycol may be employed as the solvent {See page 37). C. Apathy's mountant. Technique: 1. Thin slices of tissue are fixed for ten minutes at 37-40° C. in Solution A. 2. Transfer the material directly from the fixative to the freezing microtome and cut sections about 5/z in thickness. D 23 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 3. By means of a camel-hair brush, moistened with 50% alcohol, transfer sections from the microtome knife directly to the first of a series of dishes previously arranged as follows, in order of their use : 70% alcohol, Sudan Black solution (as above), 50% alcohol ; distilled water. 4. After immersion in 70% alcohol (in the first dish) for two minutes, stain in the Sudan Black for ten minutes, or longer if time permits. 5. Rinse in the 50% alcohol. 6. Transfer to the distilled water : mount in Apathy's medium, or, if there is sufficient time, counterstain for about three minutes in Carmalum; then rinse in distilled water before mounting. Results: Neutral fat and myelin : blue-black to black; nuclei: red. GELATINE EMBEDDING This method of embedding is employed when sections of loose friable tissues are required. Dehydration is entirely eliminated since the embedding takes place directly from water. The gela- tine which is retained during the staining processes holds the tissues together without absorbing the stain itself. Solution required: A. Gelatine 5% Gelatine . . . . . . • • 5 gi^« Phenol I % aqueous . . • • 95 ^* Warm in an oven to about 45° C. then stir until the gelatine has dissolved; raise to about 60° C. before filtering through fine calico. B. Gelatine 10% Phenol I % aqueous . . . . 90 ml. Gelatine . . . . . . . . 10 gip. Prepare exactly as for Solution A. 24 SECTION ONE C. Gelatine 15% Phenol 1% aqueous . . . . 85 ml. Gelatine . . . . . . • • 15 gm* Prepare exactly as for Solution A, except that temperature should be raised to about 75° C. before filtering. D. Gelatine 20% Gelatine . . . . . . . . 20 gm. Phenol 1% aqueous . . . . 80 ml. Prepare as for Solution A, but raise temperature to 95° C before filtering. E. Gelatine 1% Gelatine . . . . . . . . i gm. Phenol 1% aqueous . . • • 99 rnl. Dissolve by warming. Technique: 1. Small pieces of tissue, not more than 3 mm. in thickness are fixed for twenty-four hours in 5% formalin in physiological saline. 2. Wash in running water. 3. Inmierse in 5% gelatine in an incubator at 37° C. for twenty- four hours. 4. Immerse in 10% gelatine overnight in an incubator at 37° C. 5. Immerse in 15% gelatine in an incubator for several hours at 37° C. 6. Embed in 20% gelatine and leave to set. 7. Cut out blocks of tissue and immerse them in formalin for twenty-four hours. Note: The blocks may be stored indefinitely in this formalin solution if desired. 8. Rinse blocks in water and trim. 9. Freeze blocks thoroughly until they are uniformly white. 10. Allow the block to thaw somewhat until the knife cuts easily. 25 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 11. Cut sections up to 5/* in thickness and float onto slides with distilled water. 12. Drain off excess water and float sections on slides with two or three drops of 1% gelatine. 13. Drain off excess 1% gelatine and leave the slides in an oven at 37° C. until the sections are dry. 14. Immerse slide in 10% formalin for ten minutes to fix the gelatine; then stain in the usual manner with Sudan 3, or Scarlet R, Nile blue or osmic acid, or store the slides in the 10% formalin until required. LOW VISCOSITY NITROCELLULOSE (L.V.N.) For embedding tissues The following technique, using L.V.N, in place of Celloidin, has been developed by E. H. Leach and W. Chesterman, of The University Laboratory of Physiology, Oxford.* I am indebted to the authors and to the Oxford University Press for permission to print this description of the procedure. Chesterman and Leach's technique using Low Viscosity Nitro- cellulose (L.V.N.) offers advantages over the older method of em- bedding in Celloidin, in that penetration is quicker, considerably thinner sections can be cut, it is easier to use and considerably cheaper than Celloidin. With L.V.N, technique large blocks, such as half a cat's brain, can be cut at i^/u; small blocks 5x5 mm., can be cut at 5 to jju on a paraffin microtome without any special modification or attachment. L.V.N, is supplied damped with normal butyl alcohol; it is more explosive than Celloidin and it should be handled with care. When dry it will explode if hit. Exposure to sunlight should be avoided. Solutions required: Note: Solutions A and B each contain 20% of Nitrocellulose: *Q.y.M.Sc. Vol. 20, pt. 4, Dec. 1949. 26 SECTION ONE A. Absolute alcohol . . . . 210 ml. Ether . . . . . . . . 250 ml. Dibutyl phthalate . . . . 5 ml.* Mix well and add 140 gm. L.V.N, (as supplied damped with N. butyl alcohol). * NOTE In the original paper (Chesterman and Leach, 1949) and in the previous edition of this book (1953), tricresyl phosphate was stipulated: this has now been replaced by dibutyl phthalate, and I am indebted to Professor F. Bergel of The Chester Beatty Research Institute, Institute of Cancer Research, Royal Cancer Hospital, London, for calling my attention to the potential danger of handling tricresyl phosphate. Professor Bergel informs me that tricresyl-o- phosphate is highly toxic, more than 7-30 mg./kg. producing severe paralysis, and while the corresponding meta- and para-compounds show little sign of having this toxicity, the tricresyl phosphates on the market, up to the present time, always contain some of the ortho compound. References: Aldridge, W. N. (1954), Biochem.jf., 56, no. 2, 185-9, " Tricresyl Phosphates and Cholinesterase". Thompson, R. H. S. (1954), Chem. and Ind., 749, Martindale's Extra Pharmac. (1941), vol. i, 200-1. B. Absolute alcohol . . . . 210 ml. Ether . . . . . . . . 250 ml. Mix well and add 140 gm. L.V.N, (damped with N. butyl alcohol). From Solution B prepare also 5% and 10% solutions by diluting with a mixture consisting of equal parts of absolute alcohol and ether. C. Xylol . . . . . . . . 2 parts Toluol . . . . . . . . I part Beechwood creosote . . . . i part Procedure for embedding tissues: 1. Fix and dehydrate tissues as usual; then immerse in ether- absolute alcohol (equal parts) for one day. 2. Immerse in 5% L.V.N, for three to five days. 3. Transfer to 10% L.V.N, for one to two days. 4. Transfer to 20% L.V.N. (Solution B above) for one to five days. 5. Embed in Solution A. 6. Allow to harden slowly in a desiccator. In one to five days the block should be adequately hard. At this stage it should be a 27 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Stiff but easily deformable gel not altered in shape or size by shrinkage; it should be considerably less hard than a Celloidin block is usually made. 7. Plunge the block into 75% alcohol. Change the alcohol twice over a period of one to three days. 8. Trim the block, removing the hard outer rim of the L.V.N. Use 20% L.V.N, to mount it on the wood or fibre block. Harden- ing is complete in a few minutes. Dip into 75% alcohol for a few more minutes. Cutting sections: Cut the sections " dry ". If a Celloidin microtome is used the tilt of knife should be the same as that used for cutting Celloidin, but the angle the knife makes with the direction of travel should be between 25° and 45° instead of the usual 75° used for Celloidin sections ; this prevents the rolling of sections. Procedure for handling sections: 1. Collect the sections in 75% alcohol; handle and stain as usual. Dyes tend to stain L.V.N, less than Celloidin. 2. Mount sections on to a slide from a bowl of 96% alcohol. Flatten with tissue paper moistened with 96% alcohol ; press the paper with a glass rod and then remove it. Repeat this several times. 3. Treat similarly several times with equal parts of absolute alcohol and chloroform. 4. Treat similarly several times with the solution C. 5. Treat similarly several times with xylol. 6. Mount in balsam. MERCURIC CHLORIDE PRECIPITATES IN SECTIONS: METHOD FOR REMOVAL Before proceeding to the staining of sections of tissues which have been fixed in fluids containing mercuric chloride it is neces- sary to carry out the following procedure, which is essential for the removal of the deposits of mercuric chloride which would obscure the picture. 28 SECTION ONE Solutions: A. Iodine . . 0-5 gm. Alcohol 70% 100 ml. B. Sodium hyposulphite • • 7-5 gm. Alcohol 96% 100 ml. Distilled water . . 900 ml. A crystal of thymol should be added to the stock bottle. Technique: 1. Sections are mounted on slides and the paraffin wax removed with xylol. If there is any doubt as to the nature of the fixative which has been used, examine a section under the microscope: mercuric chloride, if present, will be seen as a fine brown granular deposit, more abundant in the centre of the section than on the outer edges. 2. Wash with absolute, followed by 90% alcohol. 3. Immerse for half to two minutes in Solution A. 4. Wash well with water. 5. Immerse in Solution B for half to two minutes or until the natural colour of the sections has been restored. 6. Rinse well with water. 7. Stain, dehydrate and clear in the usual manner. PARAFFIN WAX EMBEDDING AND SECTIONING {a) Dehydration Technique: 1 . Pieces of tissue are fixed and washed by any of the standard methods. 2. Immerse in 50% alcohol from twelve to twenty-four hours. 3. Transfer to 70% alcohol for the same length of time as stage 2. 4. Transfer to 90% alcohol for the same length of time. 5. Transfer to 96% alcohol for the same length of time. 6. Immerse in absolute alcohol for the same length of time. 29 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Note: Where possible use a series of corked specimen tubes for the above procedures. By occasionally shaking the tube containing the specimen the process of penetration of the alcohol is acceler- ated. Twelve hours in each change of alcohol is sufficient for small pieces, but larger pieces of tissue usually require eighteen or twenty-four hours. Hard tissues may be softened by Lendrum's technique which consists of immersing the tissues in 4% aqueous phenol for one to three days, after washing out the fixative; de- hydration is then carried out in the manner described above. Rapid dehydration of small slices of tissue: Thin slices not more than 5 mm. in thickness are immersed for half an hour in each of 50%, 70%, 90%, 96% and absolute alcohol. {h) Clearing Xylol, cedarwood oil, benzene, toluene or chloroform are the reagents most frequently used for this purpose. Xylol is the most rapid of these in displacing the absolute alcohol, but it has the disadvantage of rendering tissues brittle; therefore, if xylol is used as the clearing agent tissues must be sub- jected to it only for the minimum time necessary to displace the absolute alcohol. Cedarwood oil is slow in its action but it has the advantage of not hardening tissues even after prolonged immersion. Benzene is the best clearing agent and may be employed for the most delicate tissues : it causes the minimum shrinkage, penetrates tissues fairly rapidly and subsequently evaporates from them in the paraffin embedding bath. Toluene is also a very satisfactory clearing agent in that tissues can be subjected to it for at least twenty-four hours without risk of their undergoing shrinkage. Chloroform hardens tissues to a lesser degree than xylol but requires two or three times as long to penetrate and clear the tissue. It rapidly evaporates from the paraffin embedding bath, and it is particularly suitable for large pieces of pathological tissue. Technique: I. Small pieces of material not more than 5 mm. thick may be cleared by immersing for fifteen to thirty minutes in each of two 30 SECTION ONE changes of xylol or cedarwood oil or benzene or toluene. Larger pieces up to i cm. thick require one-and-a-half to three hours in each of two changes of the clearing agent, while bulky specimens such as whole embryos require up to six hours in each of the two changes. If at the end of the times prescribed above the specimens are not translucent or transparent they should be left in the clearing agent until they have reached that stage. (c) Embedding Technique: 1. Transfer the object from the clearing agent to a mixture con- sisting of approximately equal parts of paraffin wax and the clearing agent in a tube and place the whole in the oven set at a temperature from about 50 to 60° C. for one half to sixteen hours, depending upon the size and nature of the object; half an hour is sufficient for objects up to 3 mm. thick; an hour for 5 mm., two hours for i cm., and from eight to sixteen hours for bulky specimens such as whole embryos. 2. Transfer to pure paraffin wax in the oven from a quarter to eight hours. 3. Transfer to another bath of pure paraffin wax for the same length of time. Note: Specimens up to 3 mm. thick usually require half an hour in each of the two baths of pure paraffiua wax, while specimens 5 mm. in thickness require about an hour; and i cm. about four hours; very bulky objects about eight hours in each of the two baths of wax. Pathological material containing thrombi, emboli, etc., striated and non-striated muscle, organs containing a large amount of blood (spleen, etc.), and fibrous tissue should be subjected to im- mersion in the embedding baths for the minimum time necessary for the wax to penetrate thoroughly, as they are particularly liable to hardening and shrinkage when exposed to heat for prolonged periods. Casting the Paraffin Block I. Smear the inside of the embedding angles and the embedding base-plate very thinly with liquid paraffin ; then adjust the angles on the plate to form a mould of a suitable size. 31 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 2. Fill the mould with molten paraffin wax; then place the object in the wax and arrange it so that it is set in the right plane for sectioning. 3. When the wax block so formed is partially set, immerse it gently, while still in the mould, in cold water to ensure rapid cooling and thereby obviating crystallization of the wax and conse- quent crumbling of the block when it is mounted on the micro- tome and sections are cut. Cutting of Sections This can only be learnt by practical experience under skilled guidance in the laboratory, and it is not proposed to make any attempt to deal with the subject here since space does not permit a description of the various types of microtomes available and the technique of manipulating them. However, readers would find the chapter on section cutting in Histological TechniquCy by H. M. Carleton and E. H. Leach (published by Oxford University Press), very informative. Mounting Sections on Slides and Hydrating 1. Wet the tip of the finger slightly with glycerine albumen (Mayer) and make a smear over an area large enough to take the section in the centre of the slide. 2. Pick a section up with a needle or forceps and place it over the smear of albumen. 3. With the thumb, gently press the section on to the smear so that it is quite flat and without folds or creases, taking care not to damage the section in the process. Note: If the sections are curled up or folded, first place a drop of 1% potassium dichromate on the slide and float the section on this; then heat very gently until the section floats out flat. Blot round the edges of the section to remove excess solution; then carefully but thoroughly blot until all traces of liquid are removed. Leave the slide on a warm surface for a few minutes to drive away the last traces of water ; then proceed as follows : 4. Gently warm the slide until the paraffin wax just melts; then wash away all traces of wax with two or three changes of xylol. 5. Remove the xylol by washing the preparation thoroughly with absolute alcohol. 32 SECTION ONE 6. Wash with two changes of 90% alcohol. 7. Wash with 70% alcohol. 8. Wash with two changes of distilled water if an aqueous stain is to be used ; but if an alcoholic stain will be used staining may commence immediately after washing with 70% alcohol. 9. Proceed to stain in accordance with the staining technique it is desired to employ. Note: If the section appears opalescent when the xylol or when the absolute or the 90% alcohol is added the presence of water is indicated and it is necessary to retrace each step until the prepar- ation no longer appears opalescent when taken down to alcohol. PARAFFIN WAX - PYRIDIN TECHNIQUE A rapid method of dehydrating and clearing 1. Material is fixed and washed by the standard method. 2. If a fixative containing mercury has been used, remove mer- curial precipitate by the standard technique. 3. Immerse the tissue in two changes of Pyridin for two to eight hours in each, according to the nature and thickness of the tissue. 4. Transfer to a mixture of equal parts of molten paraffin wax and Pyridin in the embedding oven for one half to sixteen hours, depending upon the size and nature of the object. 5. Transfer to pure paraffin wax in the oven for a quarter to eight hours, depending upon the thickness and nature of the material. 6. Cast into block and cut sections in the usual manner. WATER WAX (Michrome) A very rapid and simple method of embedding tissues, obviating the use of dehydrating and clearing agents. Water wax is an amorphous water-soluble wax which sets at 56° C. to form translucent blocks similar in appearance to paraffin wax but with the complete absence of any trace of crystallization. Fresh or fixed material may be used. 33 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Reagent required: Water wax (Michrome). Technique: 1. Shake off excess water before immersing the pieces of tissue directly into a bath of water wax in the embedding oven at 55-6° C. and leave for an hour. 2. Transfer to a second bath of water wax and leave for an hour. 3. Transfer to a third bath of wax for an hour, or longer. 4. Cast the block and leave it to cool in the atmosphere or in a refrigerator. Care must be taken not to allow the block to come into contact with water. 5 . After cutting sections, float them out on water, which dissolves away the wax, and take them up on slides. 6. Stain, dehydrate and mount immediately in the usual way. Notes: (a) Fat, if present, should be dissolved out with several changes of acetone, before the tissues are immersed in the wax. {h) Blocks should be kept dry and stored in airtight containers as the wax is liable to take up moisture from the atmosphere. MISCELLANEOUS DEHYDRATING AND CLEARING REAGENTS BUTYL ALCOHOL TERTIARY For dehydrating and clearing tissues for paraffin embedding in place of ethyl alcohol and xylol. The reagent is miscible with water and with paraffin wax, and causes less shrinkage and harden- ing of tissue than does ethyl alcohol and xylol. It is also a useful substitute for ethyl alcohol for dehydrating material stained with methylene blue and other dyes which are easily extracted by ethyl alcohol. Techjiique: After fixing and washing tissues in the usual manner pass into : 1. Tertiary butyl alcohol (T.B.A.) 50% aqueous for 1-2 hours. 2. 70% Aqueous T.B.A. 2 hours to several days. 3. 85% aqueous T.B.A. for 1-2 hours. 34 SECTION ONE 4. 95% aqueous T.B.A. for 1-3 hours. 5. Pure T.B.A. for 3 changes of 4 hours in each. 6. Equal parts of Uquid paraffin and T.B.A. for 1-2 hours. 7. Infihrate in paraffin wax. CAJEPUT OIL For clearing Phis reagent will absorb small amounts of water without cloud- ing, and it is, therefore, particularly useful in wet climates as a clearing agent in place of xylol. Cajeput oil is considerably more expensive than xylol, however. CELLOSOLVE (Ethylene glycol monethyl ether) For dehydrating thin slices of tissue and sections This reagent, which is a colourless, inflammable liquid, mis- cible with water, alcohol, xylol, cedarwood oil, clove oil, and various other oils and solvents, and is also a good solvent for many stains, is coming into increasing use both for animal and plant histology, in place of ethyl alcohol : in fact many laboratories in Great Britain, at least, employ cellosolve in preference to the ethyl alcohol technique, which they now regard as obsolete. However, although some workers believe cellosolve to be superior to all other dehydrating agents as it obviates hardening and dis- tortion of most tissues, it is unsuitable for bulk material as it tends to cause distortion of protoplasmic cells owing to the rapidity of its dehydrating action. Technique : 1. Wash pieces of tissue, not more than 5 mm. thick and im- merse directly into cellosolve for half an hour. 2. Immerse in a fresh bath of cellosolve for half to one hour. 3. Immerse in a third bath of cellosolve for the same time. 4. Complete the dehydration in a fourth bath of cellosolve for an hour-and-a-half. 35 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 5. Clear in xylol for an hour. 6. Immerse in a bath of molten paraffin wax for half an hour. 7. Transfer to a second bath of paraffin wax for an hour. 8. Complete the infiltration by immersing in a third bath of wax for an hour ; then cast the block and cut sections. 9. Fix sections to slides ; then remove paraffin wax with xylol. 10. Rinse in two changes of cellosolve. 1 1 . Apply the stain ; then wash with water, or alcohol. 12. Immerse for one to three minutes in each of three changes of cellosolve. 13. Clear in xylol, benzol, clove oil or cedarwood oil. 14. Mount in balsam, cristalite or D.P.X. (Lendrum and Kirk- patrick). DIOXANE For the dehydration and clearing of tissues This reagent, which is a colourless inflammable liquid, solidi- fying at 10° C, miscible with water and paraffin wax, alcohol and xylol as well as many other organic solvents of the aliphatic and of the aromatic series, is used and preferred by many workers in place of the orthodox alcohol-xylol-cedarwood oil method for de- hydrating and clearing tissues, as the technique is simpler and quicker and it has the advantage of eliminating brittleness and shrinkage of tissues. It is, however, essential that a reliable brand of Dioxane be employed, as some makes contain relatively large amounts of water, and are therefore unsuitable for histology. Warning* — Care should be taken as Dioxane vapour is toxic: it should be used only where ventilation is abundant. Technique: I. Transfer tissues directly from the fixative to a well-stoppered specimen jar containing Dioxane with a thin layer of anhydrous calcium chloride over which is placed a circle of surgical or zinc gauze to separate the tissue from the dehydrating agent. Allow the Dioxane to act from three to twenty-four hours, depending upon the size and thickness of the tissue. 36 SECTION ONE Note: Tissues which have been treated with a fixative such as Miiller or Zenker, containing potass, dichromate, must be washed from two to twelve hours in running water, depending upon the size and the nature of the tissue, before being transferred to Diox- ane. 2. Transfer to a mixture of equal parts of paraffin wax and Diox- ane for half to one hour in a paraffin embedding oven. 3. Transfer to pure paraffin wax, allowing a somewhat longer time for impregnation than for tissues cleared by the orthodox method. Note: The Dioxane can be used several times provided the calcium chloride is changed. ETHYLENE GLYCOL A solvent, inflammable when heated, burning with an intense, almost invisible flame; which may also be employed as a de- hydrating and diflFerentiating agent for the Sudan colours, in place of acetone-alcohol, 70% or 50% alcohol; giving stable solutions, without loss of stain from the lipid particles. Solution required Technique : 1. Prepare the staining solution by heating and stirring about 0*75 gm. of the dye with 100 ml. pure anhydrous ethylene glycol to 100° C. Filter the solution while it is still hot, and again after it has been allowed to cool. 2. Cut frozen sections, from formalin fixed material, and wash them in water for about five minutes to remove excess formalin. 3. Dehydrate the sections by agitating them gently with a camel hair brush for three minutes in each of two changes of pure anhydrous ethylene glycol. 4. Transfer the sections to the staining solution for five to seven minutes, agitating them gently at intervals. 5. Differentiate by agitating the sections gently in 85% ethylene glycol in water for two to five minutes, controlling by examining under the microscope at intervals. 37 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 6. Transfer to a large dish of distilled water for three to five minutes. 7. Float sections onto slides; drain and carefully blot away- excess water. 8. Mount in glycerine jelly, Farrant or Aquamount. Note: Either ethylene or propylene glycol may be used; how- ever, the former usually costs less than the latter. Reference: Chiffelle, T. L. and Putt, F. A., (195 1), Stain Tech., 26, 51-6. PROPYL ALCOHOL, NORMAL (OR ISO) For dehydrating and clearing tissues prior to embedding Technique: 1 . Pieces of fixed tissue are placed directly into normal propyl alcohol and left therein overnight. 2. Transfer directly into a bath of paraffin wax M.P. 40° C. 3. After infiltration of the 40° C. wax, transfer to a bath of 52-54° C. paraffin wax for a few minutes; then cast the block. Note: This method prevents hardening and distortion of tissue : it is particularly recommended for scirrhous carcinoma, connec- tive tissue, tumours, etc. TERPINEOL For dehydrating sections Technique: 1. After staining and before dehydrating, wipe the slides and blot sections carefully, without allowing them to dry completely. 2. Transfer to terpineol and agitate for a few seconds. 3. Immerse in a second lot of terpineol for a few seconds. 4. Drain and wipe the slides carefully. 5. Clear with xylol, and mount as usual. Note: The destaining action of alcohol is avoided with this method, which is harmless to the vast majority of stains. Neutral Red being an exception. If desired, Cajeput Oil may be used for clearing. 38 SECTION ONE AQUEOUS MOUNTING MEDIA APATHY GUM SYRUP MOUNTANT A fairly quick-drying aqueous mountant which sets very hard and may be used in place of Farrant for fat preparations; it is, however, usually definitely acid in reaction. Hardening of the mountant may be hastened if the slides are left on warm plate. Apathy may also be used for ringing glycerine mounts. AQUAMOUNT A moderately quick-drying, transparent and colourless, aqueous mountant which, unlike Apathy and Farrant, is neutral in reaction. It takes somewhat longer to harden than does Apathy but it is unlike the latter in that it does not tend to crystallize or become excessively brittle. Aquamount is preferable to Apathy or Farrant for fat preparations. BERLEZE'S FLUID A mountant and killing fluid used in Entomology. Living specimens of Acarina and Insecta are killed by placing them directly into a drop of this fluid on the slide, but specimens which have been stored in alcohol should be washed with io% aqueous acetic acid before mounting. The fluid takes from one to two weeks or even longer to set, after which time the slides should be ringed with a waterproof cement and ringed with a layer of Canada balsam in benzene. DOETSCHMAN'S GUM CHLORAL MOUNTANT AND STAIN This is a modification, containing basic fuchsin, of Berleze's fluid, which kills, fixes, dehydrates, clears, stains and mounts a specimen in one operation. This valuable reagent is used in entomology for killing, fixing, dehydrating, clearing, staining and mounting specimens in one operation. Highly chitinized specimens should be treated with 10% potassium hydroxide from sixteen to twenty-four hours, then washed well in water before mounting directly into a drop E 39 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES of Doetschman's fluid on a slide. For permanent mounts the preparations should be placed in the oven at 37-50° C. from six to twenty-four hours. Ring the slides with a waterproof cement. Reference: MicroscopisV s Vade-Mecum nth ed., p. 207. GLYCERINE Used as a mounting medium for frozen sections. Only the purest neutral glycerine should be used. Permanent preparation may be made by painting the edges of the coverslips with melted glycerine jelly and coating the jelly, when it has set, with gold size or asphalt varnish. GLYCERINE JELLY For fat preparations, frozen sections, gelatine sections, etc. This mountant, which should be neutral in reaction, sets sufficiently hard to permit direct varnishing of the edges of the cover«lips, of mounted preparations, without prior luting. Preparations mounted in glycerine jelly will last for many years without deterioration. GLYCHROGEL MOUNTANT For Marchi-stained sections, gelatine sections, teased pre- parations, nematodes etc. A. Chrome alum . . . . . . 0-2 gm. Distilled water . . . . • . 30 ml. Dissolve by warming. B. Gelatine granules . . . . • • 3 g^i- Distilled water . . . . • • 50 ml. Glycerine . . . . . . . . 20 ml. Warm the water to 45° C. then shake in the gelatine a little at a time until dissolved and add the glycerine. Add solution A to solution B; shake thoroughly; then filter and add about o-i gm. camphor as a preservative. Keep in a well-closed bottle to prevent evaporation. Reference: Wotton, R. M. and Zwemer, R. L. (1935), Stain Tech., 10, no. i, 21-2. 40 SECTION ONE NON-AQUEOUS MOUNTING MDEIA CANADA BALSAM Strictly speaking " Canada balsam " is natural Canada balsam, a pale-yellow viscous fluid obtained from the balsam fir, indigen- ous to Canada: this fluid is useless as a histological mounting medium. "Canada balsam, dried " is the natural Canada balsam which has been dried by heat until it has become a brittle solid. *' Canada balsam in xylol " (or in benzene or other suitable solvents) is prepared by dissolving the dried natural balsam in xylol, etc. When an histologist sees the words ** Mount in Canada balsam " (or simply *' balsam ") he takes it to mean, and correctly so, that the preparation is to be mounted in Canada balsam in xylol. However, the term " Canada balsam " can be misleading to those who are new to the language and customs of histology, and such workers have been known to mount in true Canada balsam: i.e. natural Canada balsam, and with most un- satisfactory results. It is for this reason that this definition of '* Canada balsam " is given here. Although Canada balsam in xylol is still used very extensively it has a serious disadvantage in that it is prepared from a natural resin which is uncertain and variable in chemical composition, usually somewhat acid in reaction, and while it may be neutralized, will revert to an acid reaction after a time, however careful one is in handling and storing the mountant. For this reason it is better to employ a synthetic mountant of definite and unvarying chemical composition. Short descriptions of mountants of this kind are given below: they also have the advantage of being considerably less costly than Canada balsam, moreover, Canada balsam is con- siderably more costly than the synthetic mounting media des- cribed below. CLEARMOUNT Refractive index 1-515 A colourless, neutral, synthetic mountant, with a drying time approximately the same as Canada balsam in xylol. This mount- ant, which remains neutral indefinitely and does not cause the 41 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES fading of even the most delicate stains, is miscible with xylol, absolute alcohol, benzene, toluol, dioxane and many other solvents. CRISTALITE Refractive index 1-515 This synthetic mountant has all the properties of Clearmount except that it dries much more rapidly and is not miscible with alcohol. D.P.X. Refractive index 1*515 A synthetic mountant, devised by Lendrum and Kirkpatrick* on which Cristalite and probably other proprietary synthetic mountants are based. This excellent mountant, which is colour- less and neutral and does not cause fading of stains has the added advantage of being one of the least costly of all the synthetic resinous mountants: however, there is a considerable degree of shrinkage of the mountant on drying and it should therefore be applied liberally to allow for this. *y. Path. Bact.y (1939), 49, 592; (1941), 53, 441- MEEDOL BALSAM Refractive index i'5i5 A xylol miscible mountant, pale amber in colour and hardly distinguishable in appearance from Canada balsam in xylol. This is one of the least costly of all resinous synthetic mounting media. Unlike Canada balsam, but like the other synthetic mountants described in this paragraph, Meedol balsam has the advantage of remaining neutral indefinitely under normal conditions of storage and handling. FLUORMOUNT Refractive index 1*515 A synthetic, colourless, neutral, xylol miscible, non-fluorescent mountant for fluorescence microscopy. 42 SECTION ONE MICHROME MOUNTANT Refractive index 1*515 A synthetic neutral mountant miscible with alcohol and with xylol. Sections may be mounted directly from 95% alcohol if desired. Does not cause the fading of stains even after several years. This mountant, however, is largely being displaced by Clearmount. S.Q.D. BALSAM Refractive index i '5 15 An exceptionally quick-drying, synthetic, xylol miscible, synthetic mountant, similar to Meedol balsam. VENETIAN TURPENTE^E For mounting filamentous algae and other delicate material Solutions required: A. Glycerin 5%. B. Venetian turpentine 10% in abso- lute alcohol. Technique: 1. Stain the material in suitable aqueous stains. 2. Transfer to Solution A, and leave therein for several days in an open dish. 3. Wash with several changes of 95% alcohol. 4. Wash with two or three changes of absolute alcohol. 5. Transfer the material quickly from absolute alcohol to Solution B (10% Venetian turpentine in absolute alcohol), in an open dish. 6. Place the dish with contents over soda lime in a desiccator for several days until the fluid becomes viscous. 7. Mount on slides. 43 SECTION 2— ANIMAL HISTOLOGY (Normal and Pathological) 45 ALCIAN BLUE - CHLORANTINE FAST RED For selective staining of mucopolysaccharides and for morphological studies of connective tissue, cartilage and bone Solutions required: A. Ehrlich haematoxylin B. Alcian blue i% aqueous . . . . 50 ml. Acetic acid 1%, aqueous . . • • 50 ml. C. Phosphomolybdic acid 1% aqueous D. Chlorantine fast red 0-5%, aqueous Technique: 1. Fix tissues in Bouin or in 10% formalin and embed in the usual way. 2. Stain sections in Ehrlich haematoxylin for ten to fifteen minutes. 3. Blue in tap water or in lithium carbonate solution. 4. Wash in distilled water. 5. Stain for ten minutes in the Alcian blue. 6. Wash in distilled water. 7. Immerse in the phosphomolybdic acid solution for ten minutes. 8. Wash in distilled water. 9. Stain for ten minutes in the chlorantine fast red solution. 10. Wash in distilled water. 1 1 . Dehydrate ; clear in xylol and mount. Results: Nuclei are stained purplish blue. Mucin, granules of mast cells, ground substance of cartilage and some types of connective tissue 47 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES fibres, bluish green. Collagen fibres and ossein, cherry red. Cytoplasm and muscle, pale yellow. Reference: Lison, L. (1954), Stain Tech., 29, no. 3, 13 1-8. ALDEHYDE FUCHSIN (G. Gomori) For elastic tissue, mast cells, beta cells of the pancreatic islets, etc. Solutions required: A. Alcohol 70% • • • • 100 ml. Iodine crystals • • • • . . 0-5 gm. B. Sodium thiosulphate . . 075 gm Alcohol 96% • • • • 10 ml. Distilled water • • '• • . . 90 ml. C. Basic fuchsin • • • • . . 0-5 gm. Alcohol 70% • • ■ • 100 ml. Hydrochloric acid, cone. . . I ml. Paraldehyde . . • • • • I ml. Dissolve the basic fuchsin in the alcohol; to the cold solution add the acid and the paraldehyde ; shake well ; then leave to stand for about twenty-four hours until the colour of the solution has changed to violet (almost indistinguishable in appearance from gentian violet). As soon as this change has taken place the stain is ready for use. Note: The solution will keep for about four weeks at room temperature, but as the stain ages during the four weeksy longer staining times are necessary. Technique: 1. Tissues may be fixed in almost any fixative, but those con- taining dichromate are not recommended as they adversely eff"ect the clearness of the final picture. 2. Embed in paraffin wax for preference: if celloidin is used, the celloidin must be removed completely from the sections before staining as it is impervious to the stain. 3. Fix sections to slides and remove paraffin wax in the usual way. 4. Wash with absolute alcohol followed by 90% alcohol. 5. Immerse in solution A in a stoppered jar for ten minutes to one hour. 48 SECTION TWO Note: This treatment with iodine is recommended for all tissues, whether they have been fixed in mercurial fixatives or not, as it often shortens the staining time necessary and makes the shade deeper. 6. Wash well with water. 7. Immerse in solution B for about one half to two minutes until the natural colour of the section has been restored. 8. Wash well with water. 9. Stain for five minutes to two hours in a coplin jar filled with solution C. Note: Elastic fibres, five to ten minutes. Beta cells, fifteen to thirty minutes, or longer. Pituitary, thirty minutes to two hours. 10. At intervals examine the slide, after rinsing with 90% alcohol, under the microscope to ascertain the depth of staining, but taking care that the preparation is not allowed to dry. If the desired depth of staining has not been attained, the slide may be returned to the stain and rinsed with alcohol again before further examination: this process may be repeated any number of times until the desired degree of staining has been reached. 1 1 . If desired a counterstain may now be applied : haematoxylin - Orange G is best for most purposes, but for pancreas and pitui- tary, a trichrome stain of the Masson type or the Mallory- Heidenhain technique can be used to bring out all types of cells. In either case. Light Green or Fast Green, FCF should be used in place of the aniline blue as their shades contrast better with the purple of the aldehyde fuchsin. 12. Dehydrate with absolute alcohol; clear in xylol and mount. Results: The following are stained deep purple : (I) Elastic fibres of all tissues, whatever fixative has been used. (II) Mast cells, after any fixative. (Ill) The chief cells of the gastric mucosa, particularly well stained after fixation in formalin or Bouin. 49 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES (IV) Beta cells of the pancreatic islets of all species, after formalin, mercuric chloride-formalin or Bouin. Particular beauti- ful results are obtained in the islets of man and the sheep (the beta cells of the latter are particularly difficult to stain otherwise). (V) Functioning tumours of the islets are also stained select- ively. (VI) Certain basophils of the anterior pituitary, after the same fixatives as in (V). In the pituitary of the rat and of the pig the two kinds of basophils are usually quite conspicuous. Notes: {a) After fixatives containing mercury the background is pale mauve. After formalin or Bouin the background is colourless. ifi) If the stain is prepared by adding paraldehyde to Feulgen's fuchsin the resultant solution will stain the beta cells very distinctly but leave the elastic fibres unstained. Old solutions of the aldehyde fuchsin will stain elastic fibres very selectively, but leave the beta cells unstained. Reference: Gomori, G. (1950), Am. J. Clin. Path.y 20, No. 7, 665-6. ALDEHYDE FUCHSIN - HAEMATOXYLIN LIGHT GREEN - ORANGE G . CHROMOTROPE For the diflFerentiation of two types of Basophils in the Adenohypophysis of the rat and the mouse Solutions required: A. Benin's fixative with the acetic acid replaced by 0*5% trichloracetic acid. B. Lugol's Iodine C. Sodium thiosulphate 5% aqueous D. Aldehyde fuchsin Basic fuchsin 0-5 gm Absolute alcohol 60 ml. Distilled water 40 ml. Paraldehyde . . I ml. Hydrochloric acid, cone. I '5 ml. 50 SECTION TWO Note: The solution turns purple in 24 hours, and is ripe and ready for use after being kept at 20° C. for three days or in two days if kept at about 37° C. The stain deteriorates after four or five days. E. Ehrlich Haematoxylin F. Ethyl alcohol 70% .. .. .. 99-5 ml. Hydrochloric acid, cone. . . . . 0-5 ml. G. Lithium carbonate, saturated aqueous H. Orange G 2% aqueous . . . . 50 ml. Light Green SF 1% aqueous . . 20 ml. Distilled water . . . . . . 30 ml. Chromotrope 2R . . . . . . 0-5 gm. Phosphotungstic acid . . . . 0-5 gm. Glacial acetic acid . . . . . . i ml. Dissolve the phosphotungstic acid in the distilled water, then add and dissolve the chromotrope 2R followed by the acetic acid. Orange G and Light green solutions. Shake thoroughly. Note: This solution keeps indefinitely. I. Acetic Acid 0*2% Technique: 1. Fix in solution A for 24 hours. 2. Wash in running tap water for six to eight hours. 3. Dehydrate, clear and embed in paraffin wax in the usual manner. 4. Cut sections, in the horizontal plane, 3 to 4jLt in thickness. 5. Remove paraffin wax from sections with xylol. 6. Pass through the usual descending grades of alcohol to dis- tilled water. 7. Immerse in Lugol's iodine for thirty minutes. 8. Transfer to the sodium thiosulphate solution until the sections have regained their natural colour (about two minutes). 9. Rinse thoroughly in distilled water. 51 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 10. Stain in the aldehyde-fuchsin solution from two to ten minutes, taking care not to overstain, which can be avoided by checking the slides (after rinsing in 95% alcohol) at intervals under the microscope: the staining should be stopped as soon as the beta cells stand out clearly in dark purple against a colourless or faintly purple background. 11. Rinse in two changes of 95% alcohol. 12. Immerse for five to ten minutes in a third change of 95% alcohol. 13. Rinse in 70% alcohol. 14. Rinse in distilled water. 15. Stain in Ehrlich Haematoxylin for three to four minutes. 16. Rinse in distilled water. 17. Diff"erentiate by dipping three or four times in the acid alcohol solution F. 18. Blue in the lithium carbonate solution; or in running tap water for five to ten minutes. 19. Counterstain in the light green-orange-chromotrope solu- tion H for 45 seconds. 20. Rinse quickly with 0-2% acetic acid. 21. Rinse in 95% alcohol. 22. Immerse for two minutes in each of two changes of absolute alcohol. 23. Blot slides carefully. 24. Immerse in xylol for two minutes. 25. Immerse in another lot of xylol for five minutes. 26. Mount in D.P.X. or Clearmount. Results : Granulation of beta cells, selectively stained dark purple with the aldehyde-fuchsin. The cells of the pas intermedia and the Herring bodies of the neutral lobe should have little or no affinity for aldehyde-fuchsin. The delta cells are stained green, and the acidophilic granules varying shades of orange. Nuclear chromatin, purplish brown to reddish brown. Nucleoli are tinged bright 52 SECTION TWO red, by the chromotrope 2R. The non-granular cytoplasm, greyish green or unstained. Coagulated contents of the cyto- plasmic vacuoles, orange. Reference: Halmi, Nicholas, S. (1952), Stain Tech., 27, no. i, 61. ALIZARIN RED, S For calcium deposits in cartilagenous and embryonic bone Solutions required: A. Alizarin Red, S, aqueous 1% B. Polychrome Methylene Blue (Unna) Technique: Tissues are fixed in 80-90% alcohol and embedded in paraffin wax. 1. Sections are brought down to distilled water ; then stained in Solution A for five to sixty minutes, according to the material. 2. Wash with distilled water, followed by 95% alcohol at 60° C. 3. Counterstain with Solution B for one to three minutes. Results: Cartilage: intense violet. Calcium: red. Nuclei: blue. Cyto- plasm, etc. : yellow. The method is particularly suitable for pathological specimens. For bone staining in small vertebrates (Dawson's method) Solutions required: A. Potass, hydroxide 1% aqueous B. Alizarin Red, S o-i gm. Potass, hydroxide . . . . 10 gm. Distilled water i litre C. MalVs solution: Glycerin . . . . . . 20 ml. Distilled water . . • • 79 inl- Potass, hydroxide . . . . i gm. Technique: I. Whole specimens are fixed in 95% alcohol for at least three days. 53 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 2. Transfer to acetone and leave for several days to dissolve out the fats which would otherwise stain intensely and obscure the view of the bony structures. 3. Wash well with 95% alcohol; then immerse in 95% alcohol for twenty-four hours. 4. Immerse in Solution A from one to seven days, according to the size of the specimen, until the bones are clearly visible through the muscle. 5. Transfer to Solution B until the bones are stained the desired depth of colour ; this takes from one to seven days, and the solution should be changed on the fourth day. 6. Clear in Solution C until no more colour comes out. 7. Pass into a mixture of equal parts of glycerin and water, and continue through increasing strengths of glycerin. 8. Store in pure glycerin. Results: Bones are stained red ; soft tissue, transparent and unstained. Notes: If the initial clearing in potass, hydroxide solution has pro- gressed to the proper stage only the bone will be stained, but otherwise soft tissue will also be stained. The prolonged preliminary fixation in alcohol renders the tissue less liable to maceration in the potass, hydroxide solution. Objects fixed in liquids other than alcohol may be stained by this method provided they are soaked in 90% alcohol for at least three days. The best preparations are made with fish, but amphibia and mammals have also been tried with a fair degree of success, although there is not the same firm consistency about the flesh of a mammal or amphibian, prepared by this technique, as there is with that of a fish. The technique is particularly suitable for demonstrating developing bone. William's modification of Dawson's method This technique is particularly suitable for mammalian embryos and mature specimens of Urodele amphibians ; for distinguishing 54 SECTION TWO between bone and cartilage and for demonstrating the relative amount of ossification. The removal of the viscera is unnecessary in the case of museum specimens. Solutions required: A. Toluidine Blue . . . . . . 0-25 gm. Alcohol 70% . . . . . . 100 ml. Hydrochloric acid 0-5% . . 2 ml. Allow the solution to stand for twenty-four hours ; then filter and store in a tightly corked bottle. B. Potass, hydroxide 2% aqueous . . C. Alizarin Red, S . . . . . . o-ooi gm. Potass hydroxide 2% aqueous . . 100 ml. (This solution should be freshly prepared.) D. Methyl salicylate 25% in cellosolve E. Methyl salicylate 50% in cellosolve F. Methyl salicylate 75% in cellosolve Technique: 1. Wash specimens for twenty-four hours in 70% alcohol con- taining 0-2% of concentrated ammonia solution „ 2. Stain for seven days in Solution A. 3. Harden and destain for seventy- two hours in four changes of 95% alcohol. 4. Macerate for five to seven days, depending on the size of the animal, in several changes of 2% aqueous potass, hydroxide. {Note: This process is hastened by exposure to sunlight.) 5. Transfer to Solution C for about twenty-four hours when the bones should be well stained. If the specimen has been insuffi- ciently macerated the soft tissue will be slightly stained, in which case the specimen may be destained rapidly in acid alcohol (1% sulphuric acid in 95% alcohol). 6. Dehydrate by leaving the specimen in three changes of cellosolve for six hours in each. Instead of cellosolve, 50%, 80% and 90% alcohol, followed by three changes of benzol may be used for dehydration. Small embryos require less time in the dehydrating fluids. ^ 55 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 7. Clear by transferring to solutions of 25%, 50% and 75% methyl salicylate in cellosolve for twenty-four hours in each. 8. Store in methyl salicylate. Note: If glycerin is used for clearing the technique has to be modified as follows : Omit stage 6 and transfer directly from the Alizarin Red S solution into a series of 50%, 70% and 80% glycerin for twenty- four hours in each ; then store in pure glycerin. Results: Soft tissues: transparent. Osseous tissue: deep blue. Carti- lage : dark blue. Note: The relative degree of ossification and chondrogenesis which has taken place is indicated by the intensity of the stains. Bone and cartilage may be stained separately by omitting stage 2, or stage 5 for cartilage. For foetal specimens The technique is particularly suitable for mammalian embryos, for demonstrating minute bones and foetal ossification. Solutions: A. Alizarin Red, S, 1% aqueous (freshly prepared) . . . . i litre Potass, hydroxide 1% aqueous . . i ml. B. Potass, hydroxide . . . . 10 gm. Water . . . . . . . . 800 ml. Glycerin . . . . . . . . 200 ml. Note: For small specimens 5 gm. potass, hydrox- ide is sufficient. Technique: Fix in 95% alcohol for at least two weeks after making a midline abdominal incision to allow penetration of the fixative. 1. Rinse in tap water. 2. Immerse for at least four weeks in 1% potass, carbonate. 56 SECTION TWO 3. Immerse for at least ten days in 1% aqueous potass, hydrox- ide until the bones are clearly visible through the soft tissue. Note: Formalin-fixed specimens require four to six weeks in 1% potass, hydroxide. Should the tissue become too soft it may be hardened by immersing for twelve to twenty-four hours in a mixture consisting of equal volumes of glycerin, water and 95% alcohol before returning to the clearing solution. Potass, hydroxide 0*5% may be used during the last few days of the clearing. 4. Wash twenty-four hours in running tap water. 5. Stain one half to six hours, according to the size of the speci- men, in Solution A. 6. Wash for thirty minutes in running tap water. 7. Decolorize seven to fourteen days in Solution B. 8. Mount in a glass frame and dehydrate by passing slowly through alcohol-glycerin-water mixtures beginning with the pro- portions 1:2:7 and then in succession 2 : 2 : 6, 3 : 3 : 4, 4 : 4 : 2 ; and finally equal parts of alcohol and glycerin only. 9. Seal in the usual glycerin-alcohol mixture. For nervous tissues (Benda's method) Solutions required: A. Nitric acid, cone. . . . . i volume Distilled water . . . . . . 10 volumes B. Potass, dichromate 2% C. Chromic acid 1% D. Iron alum 4% E. Alizarin Red, S, saturated in abso- lute alcohol . . . . . . I ml. Distilled water . . . . . . 90 ml. F. Toluidine Blue o-i% aqueous Technique: I. Material is fixed in 90-95% alcohol for at least two days. 57 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 2. Pieces, which must not be thicker than 0-5 cm., are immersed in Solution A for twenty-four hoiirs. 3. Transfer to Solution B for twenty-four hours. 4. Transfer to Solution C for forty-eight hours ; then wash in water for twenty-four hours. 5. Dehydrate in the usual manner. 6. Clear in beechwood creosote for twenty-four hours ; then in benzol for twenty-four hours. 7. Embed in a saturated solution of paraffin wax in benzol at room temperature; then successively in saturated solutions of paraffin wax in benzol at 38° C, 42° C. and 45° C. so that pure paraffin wax only is used for the final embedding. 8. Mount sections on slides: bring down to distilled water, mordant sections on slides with Solution D for twenty-four hours, then wash thoroughly in water. 9. Stain for two hours with Solution E ; then rinse in tap water. 10. Flood slides with Solution F and warm gently until vapour is given off; or stain at room temperature for 24 hours. 11. Rinse in 1% acetic acid; then dry by blotting carefully. 12. Pass through absolute alcohol; then differentiate for about ten minutes in beechwood creosote; dry by blotting carefully, wash with xylol, and mount. Vital staining of nervous tissue in small vertebrates Solution required: AHzarin Red, S, 2% aqueous Technique: 1. Paraffin sections are brought down to distilled water by the usual method. 2. Stain twenty-four hours in 2% aqueous Alizarin Red, S. 3. Differentiate thirty to sixty seconds in distilled water to which has been added three drops 1% calcium acetate per 10 ml. 4. Dehydrate : clear and mount. Note: This is a general stain which also demonstrates Nissl bodies as well as other details. S8 SECTION TWO ALUM CARMINE - ANILINE BLUE - ORANGE G. For demonstrating the various components of the Hypophysis Solutions required: A. Cresofuchsin. B. Alum carmine (Mayer). C. Orange G . . . . . . 2 gm. Phosphomolybdic acid . . . . i gm. Distilled water . . . . . . loo ml. D. Phosphomolybdic acid 5% aqueous. E. Aniline Blue 0-2% aqueous. Technique: 1. Fix in 10% formalin; harden and dehydrate in graded alco- hols; clear in chloroform: embed in paraffin wax. 2. Sections are brought down to 70% alcohol and stained for two to twenty-four hours in Solution A. 3. Wash quickly with distilled water; then stain with Solution B for three hours, afterwards washing with distilled water. 4. Differentiate and stain the acidophil cells for five minutes with Solution C; then rinse in distilled water. 5. Immerse in Solution D for two minutes ; then blot dry. 6. Stain ten to twenty minutes with Solution E. 7. Rinse in distilled water; differentiate with 75% alcohol until no more stain comes out; then dehydrate; clear in xylol and mount. Results: Chief cells, blue to grey. Pregnancy cells, blue with small bright yellow granules; basophiles with coarse reddish blue granules. Epithelium of the pars intermedia and pars tuberalis, variable. Collagen fibre, intense blue. Glia fibres, blue-grey; axons, occasionally black. 59 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES AMMONIACAL SILVER CARBONATE For vascular reticulum, tumour cells, connective tissues around tumour, in abnormal brain tissue Solutions required: A. Pyridin, pure . . . . . . 2 volumes Glycerin, pure . . . . . . i volume B. Ammoniacal Silver Carbonate. Ammonia solution is added drop by drop to 10 ml. silver nitrate io-2% until the precipitate formed is almost redissolved, leaving a slightly opalescent solution to which is then added 10 ml. sodium car- bonate 3-1% solution and sufficient distilled water to make the volume up to 100 ml. C. Reducing solution: Sodium carbonate anhydrous . . i gm. Formalin . . . . . . . . i ml. Distilled water . . . . . . 103 ml. D. Brown gold chloride 0-2% aqueous. E. Intensifying solution: Oxalic acid 2% aqueous . . 100 ml. Formalin . . . . . . . . i ml. F. Sodium hyposulphite 10% aqueous. N.B. — All the above solutions must be kept in dark bottles. Technique: The material is fixed in 10% formalin or in Bouin and embedded in paraffin wax. 1. Bring sections down to distilled water and immerse in Solu- tion A for twenty-four hours. 2. Wash with 95% alcohol, then with distilled water. 3. Immerse in Solution B for two and a half hours at 40° C. 4. Wash with distilled water; then reduce in Solution C for five minutes, afterwards washing in tap water. 60 SECTION TWO 5. Tone for five minutes in Solution D at 30° C; then wash in tap water. 6. Intensify by immersing in Solution E for five minutes ; then rinse in tap water. N.B. — The above stages must be carried out in the darkroom. 7. Fix in Solution F. {Note: Fixation should be completed in fifteen to twenty minutes.) 8. Wash in tap water; dehydrate; clear and mount. Results: Tumour cells: reddish to greyish violet. Vascular reticulum: black. Important. — ^The tissues must not be allowed to come into contact with mercuric chloride, as even a trace will ruin the preparation. ANILINE BLUE - ACID FUCHSIN For elementary bodies in animal sections Solutions required: A. Picric acid, saturated alcoholic, . . 10 ml. Formalin . . . 25 ml. Absolute alcohol . 65 ml. Glacial acetic acid . 5 ml. B. Aniline blue, water soluble . I gm. Distilled water . 65 ml. Methyl alcohol, pure . 35 ml. Glycerin, pure . 5 ml. OxaHc acid 3% aqueous . 2 ml. C. Acid fuchsin 1% aqueous^ 100 ml. Oxalic acid 3% . . . 2 ml. Technique: I. Pieces of tissue are fixed for twenty-four hours in Solution A; washed, dehydrated, cleared and embedded in paraffin wax as usual. 61 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 2. Sections, not thicker than 5^, are fixed to slides, de-waxed and taken through descending grades of alcohol down to water as usual. 3. Stain for one half to one hour in the aniline blue (Solution B) in a stoppered staining jar. 4. Rinse well with distilled water. 5. Drain and blot carefully to remove excess water. 6. Rinse in absolute alcohol. 7. Stain for twenty minutes in the acid fuchsin (Solution C). 8. Pour off; drain and blot carefully to remove excess liquid. 9. Dehydrate with absolute alcohol; clear in xylol and mount in cristalite or in Canada balsam. Results: Elementary bodies in cells associated with the following viruses are stained scarlet : borna, zoster, rabies and pseudo rabies. ANILINE CRYSTAL VIOLET - GRAM'S IODINE For epithelial fibres Solutions required: A. AniUne crystal violet. B. Gram's iodine. C. Aniline xylol. Technique: 1. Material should be fixed in absolute alcohol and embedded in paraffin wax. 2. Sections, not more than ^fi thick, are fixed to slides and brought down to distilled water in the usual manner. 3. Stain for ten to fifteen minutes in aniline crystal violet. 4. Wash well in running water. 5. Stain with Gram's iodine for ten to thirty seconds. 6. Wash in water; drain; then blot carefully but thoroughly to remove water. 62 SECTION TWO 7. Differentiate with aniline xylol, controlling at frequent inter- vals by examination under the microscope. 8. Wash well with xylol ; mount in balsam or D.P.X. Results: Epithelial fibres are stained blue. ANILINE CRYSTAL VIOLET - LITHIUM CARMINE - IODINE For fibrin and for Gram-positive organisms in animal tissues Solutions required: A. Lithium carmine B. Crystal violet . . I gm. Aniline oil . . . . 3 ml. Absolute alcohol . . 10 ml. Dissolve and filter. C. Crystal violet 2% aqueous. D. Solution B . . 3 ml. Solution C . . 27 ml. This mixture should be prepared immediately before use. E. Gram*s iodine. F. Aniline oil . . . . I volume Xylol I volume Technique: 1. Fix material in absolute alcohol, Camoy or alcohol-formalin, and embed in paraffin wax. 2. Fix sections to slides ; de-wax and pass through descending grades of alcohol down to distilled water in the usual way. 3. Stain in the lithium carmine solution for two to five minutes. 4. Wash thoroughly in distilled water. 5. Immerse in Solution D for five to ten minutes. 63 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 6. Rinse in distilled water; drain well and blot carefully. 7. Cover with Gram's iodine solution and allow the stain to act for five to ten minutes. 8. Pour off the excess iodine solution and blot carefully with filter paper. 9. Differentiate with the aniline xylol solution until no more purple coloration comes out. 10. Drain, and blot carefully. 1 1 . Rinse with several changes of xylol. 12. Mount in balsam or in cristalite. Results: Fibrin and Gram-positive organisms are blue to blue-black while nuclei are red. ANILINE BLUE - ORANGE G (Mallory) For collagenous and reticulin fibrils ; cartilage, bone, amyloid, nuclei; fibroglia and elastin fibres Solutions required: A. Acid fuchsin J-i% aqueous. B. Aniline Blue - Orange G. Technique: Tissues are fixed in Zenker and embedded in paraffin wax, Celloidin or L.V.N. 1. Mount sections on slides and bring down to 90% alcohol; then treat with iodine in the usual way to remove mercuric deposits. 2. Bring down to distilled water and stain for one to ten minutes in Solution A; then without washing: 3. Stain for twenty minutes to one hour or longer in Aniline Blue - Orange G ; then remove excess stain with several changes of 95% alcohol. 4. Dehydrate with absolute alcohol ; clear in xylol and mount in Cristalite. 64 SECTION TWO Note: If Celloidin or L.V.N, sections are used the staining time may be shortened and 95% alcohol should be used for decolor- izing and dehydration ; terpineol for clearing. Results: Collagenous fibril, intense blue. Ground substances of cartilage, bone, mucus, amyloid : varying shades of blue. Nuclei, myoglia, neuroglia fibrils, axis cylinders, fibrin, nucleoli : red. Blood cor- puscles and myelin: yellow. Elastic fibrils: pale pink or pale yellow, or unstained; fibriloglia: red or unstained. Note: By omitting the acid fuchsin the collagenous fibres are more sharply defined. AZAN STAIN (Heidenhain) Solutions required: A. Azocarmine B .. .. 0-5 gm. Distilled water . . . . 100 ml. Glacial acetic acid . . . . i ml. Dissolve by warming ; cool and filter. B. q6% alcohol . . . . 100 ml. JLJ . Aniline oil . . o-i ml. c. 95% alcohol Glacial acetic acid . . . . 99 ml. . . I ml. D. Phosphotungstic acid Distilled water Methyl alcohol . . 5 gm. . . 75 ml. . . 25 ml. E. Aniline Blue, water soluble Orange G Glacial acetic acid . . Distilled water . 0-5 gm. . . 2 gm. . . 8 ml. 100 ml. Dissolve by warming ; cool and filter. For staining, dilute i volume of this solu- tion with 3 volumes of distilled water. 6s MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1 . Zenker-, Bouin- or Camoy-fixed tissues are stained from forty- five to sixty minutes at 55° C. in Solution A; then at room tem- perature for five to ten minutes. 2. Wash in distilled water ; then differentiate in Solution B until cytoplasm is pale pink, and nuclei are red and clear. 3. Rinse for one half to one minute in Solution C. 4. Transfer to Solution D for about one to three hours or until the connective tissue is completely decolorized ; then wash quickly in distilled water. 5. Stain for one to two hours in the diluted Solution E, exam- ining at ten- or fifteen-minute intervals to prevent over-staining. 6. Wash quickly in distilled water; then differentiate in 95% alcohol followed by absolute alcohol. 7. Clear and mount. Results: Collagen stained deep blue ; reticulum, deep blue ; chromatin, red; muscle tissue, reddish to orange; erythrocytes, red; neur- oglia, reddish ; mucin, blue. AZO CARMINE - MALLORY STAIN For Islets of Langerhans Solution required: A. Azocarmine, B, aqueous o-i% . . 100 ml. Boil for about 5 minutes; then cool and add 2 ml. glacial acetic acid. Then warm to 60 °C. and filter at that temperature. B. 90% alcohol 99 ml. -f- i nil. Aniline Oil. C. Iron alum 5% aqueous. 66 SECTION TWO Aniline Blue - Orange G (Mallory): D. Aniline blue, aqueous . . . . 0-5 gm. Orange G. . . Distilled water E. Solution D . . Distilled water . . 2 gm. 100 ml. I volume 2 to 3 volumes Technique: 1. Fix thin slices of pancreas in Bouin for eight to ten hours. 2. Wash in distilled water, dehydrate, clear and embed in paraffin wax. 3. Cut section 4/x in thickness. 4. Fix section to slides; dewax and take down through the usual grades of alcohol to distilled water. 5. Stain in solution A for about forty-five to sixty minutes at 56° c. 6. Rinse quickly in distilled water and blot very carefully. 7. Destain in solution B until acinous tissue is almost colourless and B cells show red against pink background of A cells. 8. Rinse briefly with distilled water and treat with 5% iron alum solution for 5 minutes or more. 9. Rinse again and stain two to twenty minutes in solution E until the collagenic tissue appears deep blue under the microscope. 10. Rinse and blot carefully. 11. Differentiate and dehydrate in absolute alcohol. 12. Clear in xylol and mount. Results : Cytoplasm of A cells: rich yellow orange; of B cells: bright red ; and of D cells : sky blue. Note : It is stated that it can be demonstrated that there is no gradation between A and B cells by first staining with Neutral Gentian (Bensley) decolorizing, then restaining by the above technique. Reference : Gomori, G. (1939), Anat. Rec, 74, 439-459. Cowdry, E. V.: Laboratory Technique, 3rd cd., p 167. 67 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES AZOCARMINE - HAEMATOXYLIN - ACID GREEN - ORANGE G For differential cell analysis of the rat anterior hypophysis Solutions: A. Zenker - Formol Potassium dichromate . . . . 25 gm. Mercuric chloride . . . . . . 50 gm. Ringer's solution (i.e. 0-9% saline) . . i litre. Add I mL neutral formaldehyde 40% solution per 10 ml. of the above solution immediately before using. B. Lugol's iodine C. Sodium thiosulphate 0-5% aqueous D. Delafield Haematoxylin E. Alcohol 95% Aniline oil . . 999 ml. I ml. F. Azocarmine G or B 1% aqueous . . Glacial acetic acid . . 100 ml. 4 ml. G. Glacial acetic acid . . Alcohol 90% 10 ml. 90 ml. H. Phosphotungstic acid 5% I. Acid green L extra . . Orange G . . Clove oil o-i gm. 0-5 gm. 100 ml. N.B. Use fresh stain for each batch of about twenty slides. Technique: 1. Fix in solution A for six to twelve hours. 2. Wash six to twelve hours in running water. 3. Dehydrate by immersing for thirty minutes in each of the following: 30%, 50%, 70%, 80% and 95% alcohol. 4. Immerse in absolute alcohol, two changes, for one hour in each. 5. Immerse in a mixture consisting of equal parts of absolute alcohol and cedarwood oil for one hour. 68 SECTION TWO 6. Cedarwood oil, one to sixteen hours. 7. Xylol, for fifteen minutes. 8. Infiltrate in paraffin wax 56-58° C. (four changes before finally embedding). 9. Fix sections, 4jLt in thickness, to slides and remove paraffin wax by immersing in two changes of xylol for three minutes each. 10. Immerse for three minutes in each of two changes of absolute alcohol. 11. In 95% alcohol for three minutes. 12. Distilled water for three minutes. 13. Lugol's Iodine for three minutes. 14. Sodium thiosulphate solution for three minutes when the sections should have been restored to their natural colour. 15. Stain in Delafield Haematoxylin for thirty seconds. 16. Wash in tap water for three minutes. 17. Immerse in distilled water for three minutes. 18. Immerse in 80% alcohol for three minutes. 19. Aniline alcohol (Solution E) for fifteen minutes. 20. Stain in Azocarmine for forty-five minutes. 21. Rinse in distilled water. 22. Differentiate in aniline alcohol for two to three minutes. 23. Wash in acid alcohol (solution G) for thirty to sixty seconds. 24. Immerse in phosphotungstic acid solution for one hour. 25. Dehydrate by passing through 70%, 95% and absolute alcohols (two minutes in each). 26. Counterstain in acid green - orange G solution for five minutes. 27. Clear in xylol for one minute. 28. Immerse for half an hour in each of two changes of xylol, to remove completely all traces of clove oil which would otherwise cause further decolorisation. 29. Mount in D.P.X. or Clearmount. Results: Alpha granules, purplish red. Beta cell granules, light green. Nuclear membranes are sharply defined and mitochondria are 69 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES orange red. Erythrocytes, brilliant orange. Golgi apparatus shows as negative image in both alpha and beta cells. Chromophobes show little or no cytoplasm, which is colourless to pale green. Note: A method of counting the cells is given in the original paper. Reference: Briseno-Castrejon, B. and Finerty, J. C. (1949), Stain Tech.y 24, I03-'7. BAUER - FEULGEN STAIN For Glycogen, etc. Solutions required: A. Chromic acid 4% aqueous B. Feulgen's fuchsin C. Sodium metabisulphite 10*4% aqueous . . . . . . • • 5 nil' Tap water . . . . . . • • 95 ml. Technique: 1. Fix fresh material immediately in alcohol and embed in paraffin wax. 2. Dewax sections; pass through the usual descending grades of alcohol to distilled water. 3. Immerse in the chromic acid solution for one hour. 4. Wash in running water for five to ten minutes. 5. Immerse in the Feulgen fuchsin for ten to twenty minutes. 6. Agitate the slides gently for about two minutes in each of three changes of solution C. 7. Wash in running water for ten minutes. 8. Stain nuclei, if desired, in haemalum, for two to five minutes. 9. Dehydrate as usual; clear in xylol and mount. Results: Glycogen, intense reddish-violet. Nuclei, pale mauve to navy blue. 70 SECTION TWO BIEBRICH SCARLET - ETHYL VIOLET - HAEMATOXYLIN (Cambel and Sgouris modification of Bowie's Stain) for pepsinogen granules of the body chief cells in the gastric glands Solutions required: A. Delafield Haematoxylin, aqueous B. Ethyl Violet - Biebrich Scarlet 20 % Ethyl alcohol . . C. Solution B. . . Absolute alcohol Distilled water . . 0.5 gm. 50 ml. 0-5 gm. 20 ml. 80 ml. N.B. — This solution must be freshly pre- pared each time it is required for use. D. Clove Oil . . . . . . . . I volume Toluol . . . . . . . . I volume Technique: 1. Fix in Regaud's fluid for five days in a dark phial placed in a larger amber bottle which should be wrapped round with a thick cloth and kept in a dark room. Change the Fixative daily. 2. Dehydrate with normal propyl alcohol (see page 38). 3. Transfer to paraffin wax which should be changed three times before the block is finally cast. 4. Cut sections 5 to 6/x in thickness and fix to slides as usual. 5. Remove wax with two changes of xylol. 6. Pass through absolute alcohol and the usual ascending grades of alcohols down to distilled water. 7. Stain in the Haematoxylin solution for i minute. 8. Wash and blue in tap water. 9. Remove excess water by draining and blotting very carefully. 10. Stain with solution B for ten to fifteen minutes, or longer (up to twenty-four hours). 11. Rinse briefly in distilled water, then drain and carefully remove excess water by blotting. G yi MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 12. Differentiate in solution D (clove-toluol), controlling under the microscope, for about ten to fifteen minutes. 13. Rinse with two changes of toluol. 14. Mount in permount-toluene or in Clearmount or Cristalite. Results: Zymogen granules, dark violet. Parietal cells, scarlet, and nuclei, blue. The parietal cells are distinctly contrasted from the pepsinogen cells. Reference: Cambel, P. and Sgouris, J. (1951), Stain Tech., 26, 243-6. BISMARK BROWN - METHYL GREEN For mucin, cartilage, and goblet cells in embryonic tissue, trachea and intestine Solutions required: A. Bismark brown 1% aqueous. B. Methyl green 0-5% aqueous. Technique: Tissues are fixed in Bouin or Zenker and embedded in paraffin wax. 1. Sections are brought down to distilled water; then stained five to ten minutes in Solution A. 2. Wash with 95% alcohol. 3. Stain with Solution B until the preparation appears dark green to the naked eye. 4. Dehydrate with 95% and absolute alcohol; then clear in xylol, and mount. Results: Cartilage: dark brown. Mucin: light brown. Nuclei of all cells: green. 72 SECTION TWO BIONDI - EHRLICH - HEmENHAIN STAIN For chromatin, nucleoli, mucin, etc. Solution required: Biondi - Ehrlich - Heidenhain 0-9 gm. Distilled water . . . . . . 100 ml. Dissolve by warming and stirring in a beaker. When cool add : Chloroform . . . . . . 0-25 ml. Technique: 1 . Fix tissues in saturated aqueous mercuric chloride and embed in paraffin wax in the usual manner. 2. Fix sections to slides; de-wax with xylol and pass through absolute alcohol followed by 90% and 70% alcohol. 3. Treat for the removal of mercuric precipitate by the standard technique {see page 28). 4. Immerse in the staining solution from six to twenty-four hours. 5. Rinse directly with 95% alcohol. 6. Dehydrate with absolute alcohol. 7. Clear in xylol and mount. Results: Chromatin is stained bluish green, while nucleoli are red; mucin is stained green; erythrocytes, orange. Cytoplasm and connective tissue elements are in varying shades of red. BEST'S CARMINE For glycogen Note: This method has the advantage over the Lugol's iodine technique in that fading does not occur so readily, and better staining of glycogen is obtained. The disadvantages are that the stain is less specific than in the iodine method, and the solution 73 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES deteriorates after a few weeks. If negative results are obtained the stain should be checked by staining a section known to contain glycogen. Solutions required: A. Ehrlich haematoxylin. B. Best's carmine stock solution . . lo ml. Methyl alcohol, pure . . . . 15 ml. Strong ammonia solution . . 10 ml. Note: This solution should be prepared immedi- ately before it is required for use. C. Celloidin 1% in equal volumes of absolute alcohol and ether. D. Absolute (ethyl) alcohol . . . . 80 ml. Absolute (methyl) alcohol . . 40 ml. Distilled water . . . . . . 100 ml. Technique: 1. Tissues are fixed in Bouin Fluid and embedded in Celloidin or in paraffin wax. If Celloidin sections are employed proceed as from stage 5 (below). If paraffin sections are used the procedure is as follows: 2. Float sections on the slide with 70% alcohol; flatten out; then remove excess alcohol with filter paper and blot carefully but thoroughly. 3. Remove paraffin wax with xylol in the usual manner. 4. Wash with absolute alcohol as usual. 5. Transfer the slide to a stoppered staining jar containing 1% Celloidin (Solution C, above), for fifteen minutes. 6. Transfer to a stoppered jar containing 70% alcohol, after rapidly wiping off the Celloidin from the back of the slides. This operation must be carried out quickly so that the Celloidin is not allowed to dry. Leave in the alcohol from ten to fifteen minutes. 7. Transfer to Ehrlich haematoxylin and allow the stain to act from two to ten minutes, differentiating if necessary with acid alcohol, controlling under the microscope. 74 SECTION TWO 8. Rinse in water, and without " blueing " in tap water, transfer to Best's carmine solution (formula as above) and allow the stain to act for five to ten minutes. 9. Differentiate in Solution D (above) from one to five minutes until the stain ceases to come away from the section. 10. Transfer to a mixture consisting of equal volumes of ether and absolute alcohol to dissolve out Celloidin and to dehydrate. 1 1 . Clear with xylol and mount. Results: Glycogen is stained as brilliant red granules, while nuclei are blue. BENZIDINE For brain capillaries Solution required: A. Benzidine base, pure . . . . i gm. Acetic acid 2-5% aqueous . . 200 ml. B. Sodium nitroprusside 1% aqueous C. Solution A . . . . . . 20 ml. Solution B . . . . . . 10 ml. Distilled water . . . . . . 70 ml. Mix well and filter. N.B. : This mixture should be prepared immedi- ately before use. D. Distilled water Hydrogen peroxide 20 vols. 100 ml. 1-5 ml. Technique: 1. Tissue should be fixed for one to three weeks in 10% for- malin and frozen sections, 200 to 300^ should be employed. 2. Immerse in several changes of distilled water for a total period of two hours. 75 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 3. Immerse in Solution C for half an hour at 37° C. agitating at frequent intervals. 4. Wash in several changes of distilled water. 5. Immerse in Solution D for half an hour at 37° C. agitating frequently. 6. Wash in distilled water. 7. Dehydrate by passing through ascending strengths of alcohol, beginning with 70%, in the usual manner. 8. Clear in xylol and mount. Results: Blood vessels are stained black against a background which is almost colourless. BASIC FUCHSIN - METHYLENE BLUE For demonstrating Negri L bodies in sections Solution required: Stock solutions: A. Basic fuchsin • • • • 1-5 gm- Methylene Blue . . • • • • I gm. Glycerine . . • • • • 100 ml. Methyl alcohol, pure • • • • 100 ml. B. Potassium hydroxide aqueous 0-25% .. • • • • I ml. Distilled water • • * • 99 ml. Staining solution: C. Solution A • • • • 40 ml. Solution B • • • • I ml. N.B.: This solution must be freshly prepared immediately before use. Technique: I. Pieces of fresh tissue, from the hippocampus major and cere- bellum, not more than 3 mm. in thickness are fixed in Zenker, washed, dehydrated, cleared, and embedded in paraffin wax. 76 SECTION TWO 2. Fix sections to slides ; de-wax with xylol. 3. Pass through descending grades of alcohol and treat for the removal of mercuric precipitate left by the fixative by the standard technique. 4. Place slides, section facing upwards, over the corner of a tripod. 5. Flood slide with the staining solution (Solution C, above) and heat gently with a small bunsen flame for five minutes with steam rising, taking care that the preparation does not catch fire. 6. Allow the slide to cool for a few seconds ; then wash quickly in water. 7. Decolorize and diflFerentiate in 90% alcohol until the sec- tions assume a faint violet colour. 8. Rinse quickly in 95% alcohol. 9. Dehydrate rapidly with absolute alcohol. 10. Clear in xylol and mount. Results: Negri bodies are stained deep red, while the granular inclusions are dark blue. Nucleoli are blue-black, while cytoplasm is bluish violet, and erythrocytes appear copper coloured. BASIC FUCHSIN - GENTIAN VIOLET - IODINE For bacteria in sections Solutions required: A. Basic fuchsin . . . ,. . . 0-75 gm. Alcohol absolute . . • • 30 ml. Phenol crystals . . . . . . i gm. Distilled water . . . . . . 100 ml. B. Picric acid, saturated, aqueous. C. Aniline gentian violet. 77 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES D. Gram's iodine. E. Aniline oil. . . . . . i volume Xylol . . . . . . . . I volume Technique: 1 . Pieces of tissue should be fixed in Zenker's fluid, washed in running water, dehydrated, cleared and embedded in paraffin wax in the usual manner. 2. Fix thin sections to slides, de-wax, and treat for the removal of mercuric precipitate by the standard technique. 3. Stain in the fuchsin solution for ten to thirty minutes. 4. Differentiate in formalin until the bright red colour is re- duced to pink. 5. Wash in distilled water. 6. Counterstain in the picric acid solution for a few minutes until the preparation assumes a purplish yellow colour. 7. Wash in distilled water. 8. Differentiate in 95% alcohol until the sections are red again, and yellow and red coloration begins to come out. 9. Rinse in distilled water. 10. Stain in aniline gentian violet for five to ten minutes. 1 1 . Rinse in distilled water. 12. Stain in Gram's iodine solution for one minute. 13. Drain and blot dry. 14. Differentiate with aniline-xylol until colour ceases to come out of the sections. 15. Clear with two changes of xylol and mount. Results: Gram-positive organisms are stained blue, while Gram- negative are red. Tissue, red and blue ; fibrin, deep blue. 78 SECTION TWO CARBOL ANILINE FUCHSIN For Negri bodies Solutions required: A. Basic fuchsin . 0-5 gm. Distilled water . 80 ml. Absolute alcohol . . . 20 ml. Aniline oil . . . I ml. B. Phenol . . . . Methylene Blue (Loe ffler). . I gm. Technique: 1. Fix tissues in Zenker's Fluid for twenty-four hours; wash in running water for two or three hours ; dehydrate ; clear ; embed in paraffin wax in the usual manner. 2. Sections, 4 to 5^^ in thickness, are stained from ten to thirty minutes in Solution A ; then washed with distilled water. 3. Stain with Methylene Blue (Loeffier) for fifteen to sixty seconds ; then wash with water. 4. Dehydrate and differentiate for a few seconds in absolute alcohol; then clear in xylol and mount. Results: Negri bodies are stained crimson against a blue background. CARBOL FUCHSIN (Ziehl Neelsen) For Nissl Bodies Solutions required: A. Carbol fuchsin (Ziehl Neelsen) . . 10 ml. Acetic acid 0-5% . . . . 10 ml. B. Acetic acid 1% . . . . . . 10 ml. Formalin .. .. .. .. o-i ml. 79 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Small pieces of tissue are fixed in io% formalin or in 95% alcohol or in physiological saline for at least twenty-four hours, and afterwards washed, dehydrated, cleared and embedded in paraffin wax in the usual manner. 2. Stain sections for three to four minutes with the carbol fuch- sin (Solution A). 3. Wash quickly in distilled water ; then de-stain in Solution B. 4. Wash in distilled water; then dehydrate. 5. Clear in xylol ; then mount. Results: Nissl bodies and nucleoli are stained dark red; remainder un- stained. CARBOL FUCHSIN - BORREL BLUE For Leprosy and for T.B. Solutions required: A. Carbol fuchsin (Ziehl Neelsen). B. Sulphuric acid 5%. C. Hydrochloric acid . . . . i ml. Alcohol 70% . . . . • • 99 ml. D. Borrel's Blue . . . . • • 5 ml. Distilled water . . . . . . 20 ml. Technique: 1 . Material should be fixed in saturated aqueous solution of mer- curic chloride and embedded in paraffin wax. 2. Fix sections to slides and treat them for the removal of mer- curic precipitate by the standard method [see page 28). 3. Immerse in carbol fuchsin in a staining jar for thirty minutes to an hour in the incubator at 37° C. 4. Decolorize in Solution C. 5. Decolorize in 70% alcohol (neutral) for two or three min- utes until the sections appear faintly pink to the naked eye. 80 SECTION TWO 6. Counterstain in Borrel's Blue (diluted as above: Solution D) for one or two minutes. 7. Rinse in distilled water ; drain and carefully blot away excess water. 8. Dehydrate and differentiate the Borrel Blue, controlling by examination under the microscope. 9. Clear in xylol and mount. Results: ' T.B. or leprosy, bright red; other bacteria, blue; cells and cell debris, varying shades of blue ; cell nuclei, blue. N.B, : For demonstrating leprosy, differentiation of the carbol fuchsin (stages 4 and 5) must be very carefully carried out, as this organism is more easily completely decolorized than T.B. CARBOL FUCHSIN - HAEMATOXYLIN For tubercle bacilli in mammalian tissue Solutions required: A. Alum Haematoxylin : Potash alum . . . . . . 20 gm. Haematoxylin . . . . . . i gm. Thymol . . . . . . . . i gm. Distilled water . . . . . . 400 ml. B. Carbol fuchsin (Ziehl Neelsen) i volume Distilled water . . . . • • 3 volumes Technique: Tissues are fixed in Zenker or Flemming and embedded in paraffin wax, L.V.N, or Celloidin. 1. Paraffin sections are brought down to distilled water; then stained one to five minutes in Solution A. 2. Differentiate if necessary with acid alcohol, controlling under the microscope, till nuclear detail is sharp and clearly defined; wash thoroughly in water. F 81 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 3. Stain with Solution B for five minutes, heating till steam rises ; or stain at room temperature overnight. 4. Decolorize for twenty seconds in acid alcohol; then wash thoroughly in water to which two or three drops of ammonia have been added to remove the acid. 5. Differentiate in 95% alcohol. 6. Dehydrate; clear and mount. If Celloidin or L.V.N, sections are employed, clear in terpineol or origanum oil ; blot carefully on slide and mount. Results: Tubercle bacilli, bright red. Nuclei, blue. CARBOL FUCHSESr - HAEMATOXYLIN - PICRO ACID FUCHSIN For M. leprae in sections Solutions required: A. Carbol fuchsin (Ziehl Neelsen). B. Hydrochloric acid, cone. . . 3 ml. Absolute alcohol . . • • 97 rnl* C. Potassium permanganate 1% aqueous. D. Oxalic acid 2% aqueous. E. Haematoxylin (Ehrlich). F. Picric acid, saturated, aqueous . . 50 ml. Acid fuchsin, aqueous 1% . . 10 ml. Distilled water . . . . 40 ml. Technique: 1 . Pieces of tissue are fixed for three to seven days in a mixture consisting of equal volumes of 10% formalin and absolute alcohol, and parafHn sections are employed. 2. Stain sections in the carbol fuchsin solution in a stoppered staining jar for three or four days. 82 I SECTION TWO 3. Immerse in 10% formalin, of a slightly acid reaction, for five minutes. 4. Immerse in the acid alcohol for five minutes. 5. Flood the preparation w^ith potassium permanganate and allow the reagent to act until the sections turn brown (this usually takes from two to five minutes). 6. Immerse in the oxalic acid for one minute. 7. Stain with Ehrlich haematoxylin solution for two minutes ; then blue in tap water or in saturated lithium carbonate solution aqueous. 8. Stain in picro-acid fuchsin for two to five minutes; then without washing: 9. Dehydrate, clear and mount. Results: M. leprae, dark blue. Connective tissue fibres, red. Muscle, yellow. Nuclei, brown. CARBOL FUCHSIN - IODINE - HAEMATOXYLIN ORANGE G For demonstrating leprosy organisms together with neuro- keratin of the myelin sheath Solutions required: A. Lugol's iodine. B. Carbol fuchsin (Ziehl Neelsen). C. Absolute alcohol . . . . . . 35 ml. Distilled water . . . . . . 65 ml. Hydrochloric acid concentrated. . 0-5 ml. D. Ehrlich's haematoxylin. E. Strong ammonia solutioix (sp. gr. o-88o) . . . . . . . . I ml. Distilled water . . . . • • 99 nil* F. Orange G. aqueous 1%. 83 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Pieces of tissue are fixed in Zenker; washed; then trans- ferred to a mixture of Lugol's iodine and 80% alcohol (equal vol- umes of each) for six to twenty-four hours. 2. Transfer to 80% alcohol for twelve to twenty-four hours. 3. Immerse in 95% alcohol for two to six hours. 4. Transfer to a mixture consisting of equal volumes of absolute alcohol and xylol, for half an hour. 5. Immerse in xylol for half an hour. 6. Immerse in two changes of paraffin wax before casting the block and finally sectioning. 7. Fix sections to slides and remove wax with xylol. 8. Pass through absolute, 90% and 70% alcohol. 9. Stain for half an hour in carbol fuchsin (Solution B). 10. Rinse in distilled water. 11. Partially differentiate with the acid alcohol (Solution C, above). 12. Rinse well with distilled water. 13. Stain for one to two minutes with Ehrlich haematoxylin solution. 14. Differentiate in the acid alcohol (Solution C). 15. Rinse well with distilled water. 16. Immerse in the ammonia solution (Solution E above) for a few seconds. 17. Rinse well with distilled water. 18. Stain with the Orange G solution for two to three minutes. 19. Dehydrate rapidly with two changes of acetone. 20. Clear in xylol and mount. Results: Leprosy organisms and neurokeratin are stained red, while nuclei are blue and cytoplasm is yellow. 84 SECTION TWO CARBOL FUCHSIN - METHYL GREEN For demonstrating hyaline substance Solutions required: A. Carbol fuchsin (Ziehl Neelsen) . . 5 ml. Distilled water . . . . • • 45 rnl. B. Methyl Green 1% in 5% acetic acid. Technique: 1. Material which has been fixed in any of the standard fixatives is embedded in paraffin wax. 2. Fix sections to slides : then bring down to distilled water as usual. 3. Stain in Solution A for fifteen to forty-five minutes. 4. Wash with distilled water; drain off excess; then blot care- fully. 5. Dehydrate rapidly with absolute alcohol. 6. Differentiate and counterstain in Solution B for two or three minutes. 7. Wash quickly with absolute alcohol. 8. Clear in xylol and mount. Results: Hyaline substance is stained bright red while nuclei are light green. CARBOL THIONIN - PICRIC ACID (Schmorl) For demonstrating bone canaliculi Solutions required: A. Decalcifying solution: Formalin 10% . . . . . . 100 ml. Nitric acid, cone. . . . . 15 ml. B. Carbol thionin (Nicolle). C. Picric acid 1% aqueous. 85 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Formalin-fixed specimens are placed in a large volume of Solution A, which is changed once or twice a day. The time required for complete decalcification will depend, of course, on the thickness and the nature of the specimen. The bones of young animals usually take from twenty-four to forty-eight hours, but in other cases as long as a week may be necessary. Decalcification is complete when the bone has become flexible and can easily be punctured with a needle. 2. Make Celloidin or frozen sections. 3. Rinse in water for ten minutes. 4. Stain with Solution B for 10 minutes; then rinse in distilled water. 5. Immerse in Solution C for half to one minute; then wash in water. 6. Differentiate with 70% alcohol for five to ten minutes until the stain ceases to come out of the sections. 7. Dehydrate with 96% alcohol; clear in origanum oil; then mount in balsam. Results: Ground substance, yellow to brown; bone canaliculi, dark brown to black; cells, red; ground substance of cartilage, brilliant purple. Note: Carbol thionin (NicoUe) deteriorates after a few weeks and this stain is therefore best when freshly prepared. CARMINE - METHYLENE BLUE (Schultz-Schmitz Stain) For demonstrating sodium urate in animal tissue Solutions required: A. Distilled water . . . . . . 64 ml. Lithium carbonate . . . . 0-5 gm. Carmine . . . . . . . . i gm. Ammonium chloride . . . . 2 gm. Boil for a few minutes ; allow to cool ; then make up to the original volume and add : Strong ammonia solution . . 6 ml. Filter before use. 86 SECTION TWO B. Solution A 15 ml. Pure methyl alcohol 12-5 ml. Strong ammonia solution (sp. gr. o-88o) 2 ml. Distilled water 5.5 ml. C. Methylene Blue i% in absolute alcohol. D. Picric acid, saturated aqueous . . 27 ml. Sodium sulphate saturated aqueous . . . . • • 3 nil. Technique: 1. Fix thin slices of the material in absolute alcohol. 2. Immerse for one-and-a-half to two hours in each of three changes of acetone. 3. Transfer to a mixture of equal volumes of acetone and benzol for half an hour. 4. Immerse in pure benzol for one half to one hour ; then embed in paraffin wax. 5. Fix sections to slides and de-wax with xylol. 6. Pass through absolute, followed by 90% alcohol. 7. Immerse in Solution B in a grooved staining jar for five minutes, rocking gently, but continuously, during the period of staining. 8. Rinse thoroughly with absolute alcohol. 9. Stain for half a minute in the methylene blue solution. 10. Rinse with absolute alcohol. 11. Stain for fifteen to thirty seconds in Solution D keeping the slides in motion by rocking. 12. Dehydrate thoroughly with absolute alcohol; clear in xylol and mount in balsam or cristalite. Results: Nuclei are stained greyish blue, while cytoplasm is yellowish; uric acid crystals are deep greenish blue; monosodium urate, brilliant green. H 87 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES CELESTIN BLUE - CHROMOTROPE 2R (Lendrum) A substitute for haematoxylin-eosin, for simple diagnostic or photographic purposes, emphasizing the staining of collagen and reticulum Solutions required: A. Celestin blue (as solution B, page 89) B. Alcohol 70% 100 ml. Hydrochloric acid, cone. . . . . 2 ml. C. Phosphomolybdic acid 1% D. Chromo trope 2R 1% in absolute alcohol Technique: 1. Fix tissues in Zenker or Bouin exactly as described on page 89, stage i). 2. Stain section in the celestin blue solution for fifteen minutes. 3. Remove any cytoplasmic staining with solution B. 4. Wash with water for one minute. 5. Mordant with 1% phosphomolybdic acid solution for one to two minutes. 6. Wash well with water. 7. Dehydrate; then stain for two minutes in the chromotrope 2R solution. 8. Dehydrate; clear in xylol, and mount in D.P.X. Results: Nuclei, bluish purple. Cytoplasm, pink; collagenous elements, bright red. Note: For photographing this stain the best filters to use are those giving a spectral transmission of 5,600 to 6,000 A.U. Reference: Lendrum, A. C. (1935), J- Path, and Bad., 40, 415-6, " Celestin blue as a nuclear stain ". 88 SECTION TWO CELESTIN BLUE - ORCEIN - LIGHT GREEN (Lendrum) For breast carcinoma and skin lesions Solutions required: A. Rubens - Duval Orcein: Orcein Alcohol 70% Nitric acid, cone. o-i gm. 100 ml. 2 ml. B. Celestin Blue Iron alum 5% . 0-5 gm. 100 ml. . . 2 ml. . . 14 ml. . . 2 volumes . . I volume IS. . . 100 ml. . . 2 ml. (prepared with cold distilled water) Shake the dye with the iron alum solution in a flask; then boil for three minutes. Allow to cool ; filter ; then add Sulphuric acid, concentrated Glycerine C. Eosin yellowish aqueous 1% Gallic acid 0-5% aqueous D. Phosphomolybdic acid 1% aqueous. E. Masson^s Light Green: Light green 2% aqueous . . Glacial acetic acid .... Technique: 1. Fix tissues in Zenker or Bouin; if the latter is used then picric acid must be removed by washing de-waxed and dehydrated sections on slides with saturated lithium carbonate solution. If Zenker is employed ; then mercuric precipitate must be removed after fixation by the standard technique. 2. Wash; dehydrate; clear; embed in paraffin wax as usual. 3. Fix sections to slides ; remove paraffin wax with xylol. 4. Pass through descending grades of alcohol down to distilled water in the usual manner. 5. Stain for one half to two hours in Solution A, in a stoppered grooved staining jar in the incubator or for twenty-four hours at room temperature. 89 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 6. Rinse well with distilled water. 7. Stain the nuclei with celestin blue (Solution B) for fifteen minutes. 8. Wash with running water for twenty minutes. 9. Stain muscle and epidermis for two minutes with eosin (Solution C). 10. Decolorize the collagen somewhat in water or in 30% alcohol. 11. Immerse in phosphomolybdic acid for two minutes. 12. Wash well in distilled water. 13. Stain collagen with the light green solution. 14. Dehydrate rapidly. 15. Clear in xylol; mount in D.P.X. Results: Elastin, light brown. Nuclei, bluish purple. Muscle and epider- mis, red. Collagen, green. From J. Path, and Bad., 40, 415-6, 1935 Lendrum, A. C. " Celestin blue as a nuclear stain." CHLORAZOL BLACK A general-purpose stain, which can be used for whole tissues as well as for sections. The stain requires no mordant or differenti- ation, and it may be employed in aqueous or alcoholic solution. A saturated solution in 70% alcohol stains ordinary sections in fif- teen to thirty minutes ; the stain does not fade. The stain is particularly suitable for staining embryo ; kidney ; intestine for demonstrating epithelial cells ; chromatin ; nucleoli ; muscle fibres. Solution required: Chlorazol Black, saturated in 70% alcohol. Technique: 1. Tissues should be fixed in Zenker and embedded in paraffin wax. 2. Bring sections down to 70% alcohol and remove mercuric precipitate in the usual manner. 90 <( SECTION TWO 3. Stain in a freshly prepared, unfiltered, saturated solution of Chlorazol Black in 70% alcohol for five to ten minutes. 4. Drain off excess dye ; dehydrate ; clear in xylol and mount. Results: Embryo, epithelial cell tissues, outlined in black. Chromatin, black. Nuclei, black. Muscle fibres, intense black. Lympho- cytes, intense black. Blood cells, yellowish green. Cytoplasm, greenish grey. Kidney and intestine, varying shades of green, grey and black. Blood cells, light green. Nuclei and chromosomes are stained black; cytoplasm and secreted products grey; chitin, green; glycogen, red. Notes: (a) Benzl alcohol may also be used as a solvent, in which case results are somewhat different. {h) If it is desired to differentiate chlorazal black, dilute Milton " (a proprietary antiseptic) may be used for the purpose. {c) The stain may be incorporated with Lactophenol. CONGO RED For Amyloid in tissues Solutions required: A. Congo Red 1% in distilled water. B. Lithium carbonate saturated aqueous C. Delafield or Ehrlich haematoxylin. Technique: 1. Formalin or alcohol-fixed material may be embedded in Celloidin or in paraffin wax, or frozen sections may be employed. 2. Sections are mounted on slides and brought down to distilled water as usual. 3. Stain in the Congo Red solution for ten to thirty minutes. 4. Immerse in the lithium carbonate solution for fifteen seconds. 5. Decolorize in 80% alcohol until stain ceases to come away in clouds. 91 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 6. Wash in running water for fifteen minutes ; then stain with Ehrlich or Delafield haematoxylin for five to ten minutes. 7. Wash in water; dehydrate in the usual manner; clear in xylol and mount. Note: If Celloidin sections are used dehydration should be carried out with Isopropyl alcohol in place of ethyl alcohol. Results: Amyloid, red; nuclei, blue. CONGO RED - ANILINE BLUE - ORANGE G For elastic fibres Solutions required: A. Aluminium chloride, 2% aqueous. B. Congo Red . . . . . . 2 gm. Sodium citrate . . . . . . 2-5 gm. Glycerin . . . . . . . . i ml. Distilled water . . . . • • 97 i^il- C. Aniline Blue, aqueous . . . . 1-5 gm. Orange G . . . . . . 2-25 gm. Resorcinol . . . . • • 3 gm- Phosphomolybdic acid 1% aqueous 100 ml. Technique: Tissues should be fixed in 10% formalin, and frozen sections should be employed. 1. Wash sections in water; then immerse them in Solution A for ten minutes. 2. Wash with water and drain; then stain in the Congo Red solution for ten minutes. 3. Wash with tap water ; then plunge the slide into a dish of tap water and agitate it there for ten seconds. 4. Wash again with tap water ; then stain from five to ten min- utes in the Aniline Blue-Orange G solution (Solution C above). 5. Rinse carefully in tap water; drain well and blot. 92 SECTION TWO 6. Dehydrate in absolute alcohol ; clear in origanum oil ; wash in xylol and mount. Results: Elastic fibres, bright red ; fibrin, dark blue. CONGO RED - EHRLICH HAEMATOXYLIN For eleidin and keratohyalin Solutions required: A. Congo Red 0-05% aqueous. B. Ehrlich haematoxylin. Technique: 1. Material should be fixed in absolute alcohol and embedded in paraffin wax. 2. Sections not more than 5^ thick are mounted on slides and brought down to distilled water as usual. 3. Stain for five to ten minutes in the Congo Red solution. 4. Rinse in distilled water; then stain for five to ten minutes in Ehrlich haematoxylin. 5. Blue in tap water in the usual manner. 6. Dehydrate; clear in xylol and mount. Results: Eleidin is stained red, while nuclei and keratohyalin are blue. CRESYLFAST VIOLET - TOLUIDINE BLUE - THIONIN (EHRLICH) A non-fading tri-basic stain for nerve cells and Nissl granules, in normal and pathological tissues Solutions required: A. Cresylfast violet, CNS . . . . 2 gm. Toluidine blue . . . . . . i gm. Thionin (Ehrlich) 0-5 gm. Ethyl Alcohol 30% 200 ml. 93 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES B. Distilled water . . . . . . 200 ml. Sulphuric or nitric acid, cone. . . 0-5 ml. Technique: 1. Formalin fixed material is embedded in paraffin wax, and sections, 4/x in thickness are fixed to slides with glycerin albumen. 2. Remove wax with xylol. 3. Rinse with absolute alcohol. 4. Pass through 95% alcohol. 5. Pass through 80% alcohol. 6. Immerse slides for five to ten seconds in the staining solution at 80-90° C. 7. Differentiate for one second in solution B. 8. Dip and agitate slides in a beaker of cold distilled water for one second. 9. Differentiate further in 80% and 95% alcohol for one to two seconds in each. 10. Immerse in 80% alcohol for one second. 1 1 . Dip and agitate the slides in the still warm solution A for one to two seconds. 12. Return to 80% alcohol for one second. 13. Repeat steps 11 and 12. 14. Rinse in distilled water. 15. Dehydrate by immersing for one second in each of 80%, 95% and absolute alcohol. 16. Immerse in xylol for one minute. 17. Immerse in a fresh lot of xylol for three minutes. 18. Mount and examine. Results: Neurons stand out distinctly against a pale background, and can be followed for a considerable distance. The cytons are stained dark purple emphasizing the blue tint, while the dendrite and axon processes and endings present a somewhat lighter shade, bluish to reddish. Granules in the cell body as well as in the protoplasm 94 SECTION TWO processes appear purple or reddish. Nuclei and nucleoli are well differentiated. Reference: Spoerri, Rosette (1948), Stain Tech., 23, 133-5. DAHLIA ACETIC (Ehrlich) For mast cell granules in sections Solution required: Distilled water . . . . . . 100 ml. Absolute alcohol . . . . . . 50 ml. Glacial acetic acid . . . . 12-5 ml. Dahlia . . . . . . . . 10 gm. Dissolve by heating in a flask, lightly plugged with cotton-wool, on a water bath. Allow to cool ; then filter. Technique: 1. Fix tissues in absolute alcohol and embed in Celloidin. 2. Immerse sections in the staining solution for twelve hours. 3. Differentiate in 95% alcohol. 4. Clear in origanum oil and mount in balsam or in cristalite. Results: Granules of mast cells are stained reddish violet. DOPA REAGENT For melanoblasts * Solutions required: A. Dopa reagent (3 : 4-dihydroxy- phenylalanin) . . . . 0-2 gm. Cold distilled water . . . . 200 ml. Note: This solution,^ which deteriorates fairly rapidly at normal temperatures, should be kept in a refrigerator. When the solution turns dark red it is useless and should be discarded. B. Buffer tablet pH 7-4 . . . . i tablet Distilled water (cold) . . . . 100 ml. 95 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES C. Solution A . . . . • • 25 ml. Solution B . . . . . . 8 ml. Note: This solution must be freshly prepared as required. Technique: Tissues should be fixed not longer than two to three hours in 5% formalin, or frozen sections of fresh material may be employed. 1. Rinse with distilled water from four to five seconds; then immerse in Solution C for three to four hours, controlling under the microscope at intervals, until melanoblasts are stained black. Note: This solution is likely to overstain if it becomes sepia brown. 2. Wash with distilled water; dehydrate; clear; mount. Result: Melanoblasts, black. Note: After dehydration (in Stage 2) 1% crystal violet in abso- lute alcohol may be employed as a counterstain, if desired. ELASTIN STAIN (Weigert) Any fixative may be used except Susa, and tissues may be em- bedded in paraffin wax or in Celloidin or in L.V.N. Preparation of the Staining Solution: Triturate i gm. of Weigert elastin stain and 5 gm. clean, dry silver sand with 100 ml. absolute alcohol and 2 ml. pure hydro- chloric acid until all the stain has gone into solution; then filter. Note: The staining solution deteriorates after two or three weeks. The nuclei may be stained with Orth's lithium carmine prior to the following procedure if no other counterstain is desired. Technique: I . Sections are brought down to 90% alcohol and stained one half to twelve hours according to depth of staining desired. The slides 96 SECTION TWO should be stained in a jar or in a Petri dish, sections face down- wards, to prevent a deposit forming on the sections. 2. Wash off excess stain with 95% alcohol, and if necessary differentiate in acid alcohol for a few minutes. 3. Wash quickly with 70% alcohol ; then thoroughly with water. 4 Counterstain with Van Gieson, Ehrlich haematoxylin or Safranin for about five minutes. 5. Differentiate, if necessary, in 95% alcohol. 6. Dehydrate ; clear in xylol and mount. Note: If Celloidin or L.V.N, sections are used clear in ori- ganum oil or in terpineol after 95% alcohol. Results: Elastic fibres, dark blue or black. Nuclei, brilliant red (if Orth's carmine is used) or bluish black (with haematoxylin). Collagen, pink to red ; other tissue elements, yellow (if Van Gieson is used). ELASTIN STAIN (Sheridan) This stain has an advantage over Weigert's elastin stain in that the solution may be kept for reasonably long periods without deterioration. The staining procedure is the same as for Weigert's elastin stain. Results: Elastic fibres are stained green to greenish black. ELASTIN - TRICHROME STAIN For the demonstration of elastic, smooth muscle and collagenic fibres with equal clarity, particularly in the walls of hlood vessels Solutions required: A. Weigert's elastin stain Weigert's elastin stain powder . . i gm. Hydrochloric acid, cone, pure . . 2 ml. Absolute alcohol . . . . 100 ml. 97 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Dissolve the stain by boiling for two minutes in a flask, plugged lightly with cotton-wool, on a water bath. Allow to cool; then filter; make the volume up to 100 ml. with absolute alcohol; then add the acid. Alternatively the solution may be prepared as described on p. 96. Note: This solution deteriorates after three or four weeks. B. Ehrlich haematoxylin. C. Ponceau-acid fuchsin (Masson): Acid fuchsin . . . . . . 0-3 gm. Ponceau de xylidine . . . . 0-7 gm. Distilled water . . . . . . 100 ml. Glacial acetic acid . . . . i ml. D. Phosphotungstic acid 3% aqueous. E. Light Green 1% aqueous. Technique: 1. Paraffin sections are mounted on slides and brought down to distilled water in the usual manner ; then immersed in Weigert's elastin stain in a staining jar for one hour. 2. Wash rapidly in acid alcohol; then dehydrate and differ- entiate in absolute alcohol until the sections appear only faintly red. 3. Immerse in 70% alcohol, followed by distilled water. 4. Stain in Ehrlich haematoxylin for eight to ten minutes ; then differentiate in water for five minutes. 5. Stain in Ponceau-acid fuchsin for five minutes. 6. Wash thoroughly in 3% phosphotungstic acid ; then immerse in the phosphotungstic acid for ten minutes. 7. Wash thoroughly in distilled water; then stain with Light Green for two to five minutes; then without washing: 8. Flood the preparation with 1% acetic acid and allow it to act for three minutes ; pour off excess ; then without washing : 9. Dehydrate; clear; mount in Damar xylol. q8 SECTION TWO Results: Elastic tissue stained blue-black ; smooth muscle, red ; collagen green. EOSIN AZUR 2 - HAEMATOXYLIN (Maximow) For demonstration of inflammatory changes in haemo- poietic tissues Solutions required: A. Azur 2 eosin .. .. .. o-i gm. Distilled water . . . . . . loo ml. Heat to boiling point, then allow to cool. B. Solution A (as above) . . . . lo ml. Distilled water . . . . . • 50 ml. C. Ehrlich haematoxylin. Technique: 1 . Formalin-fixed material (sections or smears) are stained from five to ten minutes with Ehrlich haematoxylin. 2. Pour off excess stain ; immerse the preparation in tap water until it appears blue to the naked eye ; then wash thoroughly with distilled water and drain well. 3. Stain from eighteen to twenty-four hours in Solution B (as above) in a staining jar. If a staining jar is not available, place the slide, resting face downwards, on two pieces of thin glass rod, so that any precipitate formed is not deposited on the preparation. 4. Differentiate in 95% alcohol until dense blue clouds cease to come away from the preparation, and the red corpuscles and collagen are pink. 5. Immerse the preparation in three changes of absolute alcohol, followed by two changes of xylol ; then mount. Results: Cartilage stained purple; basophil leucocytes and mast cell granules, purple to violet; nuclei, blue; erythrocytes, pink; cytoplasm, pink to blue ; eosinophil granules and secretion gran- ules, pink. 99 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES ETHYL VIOLET - BIEBRICH SCARLET (Bowie) For pepsinogen granules in gastric mucosa Solutions required: A. Stock solution. Ethyl violet-Biebrich scarlet i% in 20% alcohol. B. Staining solution. Stock solution (as above) . . 0-5 ml. Alcohol 20% . . . . . . 100 ml. Technique: 1. Tissues should be fixed in Regaud's fluid, washed in running water, dehydrated, cleared and embedded in paraffin wax in the usual manner. 2. Fix sections to slides, dewax and pass through descending grades of alcohol down to distilled water. 3. Stain for twenty-four hours in Solution B in a covered staining jar. 4. Drain and wipe oflF excess liquid. 5. Differentiate for about half an hour with a mixture con- sisting of equal volumes of clove oil and xylol, controlling by examination under the microscope at intervals. ^ 6. Wash well with several changes of xylol ; mount in cristalite. Results: Pepsinogen of the pepsin-forming cells is stained violet, while parietal cells are red. FEULGEN FUCHSIN For chromatin in animal cells Solutions required: A. N/i hydrochloric acid. 100 SECTION TWO B. Distilled water . . . . . . 200 ml. Boil; allow to cool to about 70° C; then add: Basic fuchsin . . . . . . i gm. When dissolved, raise to boiling point ; then allow to cool to 50° C. and add: 2 ml. pure hydrochloric acid cone, and 2 gm. potass, metabisulphite. Allow to stand twenty-four hours then add i gm. decolorizing carbon; shake well; filter. The solu- tion if properly prepared will be colourless. C. Pure hydrochloric acid, cone. . . 2 ml. Potass, metabisulphite . . . . 2 gm. Distilled water . . . . . . 200 ml. D. Light green 0-25% in 90% alcohol. Technique: Tissues are fixed in Zenker or Helly for six to twelve hours ; then washed in running water for twelve to twenty-four hours. They should not be treated with iodine-alcohol. 1. Sections are transferred from distilled water to ajar of Solu- tion A at 60° C. for five to fifteen minutes; then rinsed in distilled water. 2. Transfer for one to one and a half hours to a jar of Solution B at room temperature. 3. Transfer for two to three minutes in each of three successive lots of Solution C ; then wash thoroughly in distilled water. 4. Counterstain for about two to five minutes in Solution D. 5. Dehydrate; clear and mount. Results: Chromatin, purple. All other elements, transparent green. lOI MEDICAL AND BIOLOGICAL STAINING TECHNIQUES FONTANA STAIN For argentaffine granules Solutions required: A. Silver oxide (Fontana). B. Sodium thiosulphate 5% aqueous. Technique: 1. Tissues are fixed in 10% neutral formalin, washed, dehy- drated in alcohol, cleared in cedarwood oil, and embedded in paraffin wax as usual. 2. Fix sections to slides, bring down to distilled water and wash thoroughly in two or three changes of neutral, freshly distilled water. 3. Immerse in the silver oxide (Fontana) solution for twelve to twenty-four hours in the dark in a covered, scrupulously clean vessel. 4. Wash in neutral, freshly distilled water for one minute. 5. Immerse for one minute in the sodium thiosulphate. 6. Immerse in tap water for ten minutes. 7. Counterstain, if desired in carmalum. 8. Dehydrate; clear in xylol and mount. Results: Argentaffine granules, black. FONTANA STAIN - SILVER NITRATE For reticular and collagen fibres Solutions required: A. Strong ammonia solution (sp. gr. o-88o) . . . . . . . . 10 ml. Distilled water . . . . . . 90 ml. B. Potassium permanganate 0-5% aqueous. C. Oxalic acid 1-5%. 102 SECTION TWO D. Silver nitrate 5% aqueous. E. Fontana stain (silver oxide solution) F. Aniline oil . . . . . . . . i volume Xylol . . . . . . . . I volume Technique: 1. Frozen sections not thicker than lo/^ are fixed in 10% for- malin and afterwards washed in three changes of water for fifteen minutes in each. 2. Immerse in Solution A at 60° C. for fifteen minutes, in an oven. 3 . Rinse well in three changes of distilled water. 4. Immerse in the potassium permanganate for three or four minutes. 5. Rinse with distilled water for about ten or twenty seconds. 6. Decolorize with the oxalic acid solution until the brown colour just disappears ; then wash well in distilled water. 7. Immerse in silver nitrate solution in the dark for an hour. 8. Wash well with two changes of distilled water in the dark. 9. Immerse in Fontana's stain for fifteen minutes at 60° C. in the dark. 10. Wash rapidly in three changes of distilled water. 11. Immerse in 30% formalin for two or three minutes at 60° C. 12. Wash thoroughly in running tap water; then transfer to slides. 13. Blot away excess water. 14. Dehydrate with two changes of absolute alcohol. 15. Clear in the aniline-xylol (Solution F, above). 16. Wash with xylol; mount in dammar xylo Results: . Reticulum is stained black, while collagen is brown. N.B. : Sections must be handled with glass needles throughout this technique, as contact with metal instruments will ruin the preparations. I 103 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES GALLOCYANIN - ORCEIN - ACID ALIZARIN BLUE - ALIZARIN VmiDINE A general stain for animal tissues Solutions required: A. Gallocyanin .. .. .. o-i gm. Chrome alum 5% aqueous . . 100 ml. Boil for ten minutes. Allow to cool ; then make up the volume to 100 ml., filter and add five or six drops of formalin. B. Orcein 0-5% in 70% alcohol . . 99 ml. Hydrochloric acid cone. . . i ml. C. Acid Alizarin Blue . . • • 5 gn^- Aluminium sulphate 10% aqueous 100 ml. Boil for ten minutes. Cool and filter. Make up the volume to 100 ml. with distilled water and add five or six drops of formalin. D. Phosphomolybdic acid 5%. E. Alizarin viridin . . . . . . o-2 gm. Buffer solution pH 5-8 . . 100 ml. Technique: 1. Fix tissues in 10% formalin and embed in paraffin wax in the usual manner. 2. Fix sections to slides and take down to distilled water as usual. 3. Stain nuclei intensely by immersing the slides in the gallo- cyanin solution in a staining jar, examining the preparations under the microscope at intervals over a period of twenty-four hours, to ascertain the depth of staining. 4. Wash with two changes of distilled water. 5. Stain elastic fibres in the orcein solution for ten minutes to half an hour in a grooved, covered staining jar. 6. Wash well with distilled water. 7. Stain muscle in the acid alizarin blue solution for seven minutes. 104 SECTION TWO 8. Wash with distilled water. 9. Differentiate in the phosphomolybdic acid solution for about thirty minutes, controlling by examination under the micro- scope at intervals. 10. Wash with two changes of distilled water. 11. Stain collagen in the alizarin viridin for seven minutes. 12. Drain and blot thoroughly but carefully. 13. Rinse with 96% alcohol; followed by carbol xylol. 14. Wash well with two or three changes of xylol, and mount. Results: Nuclei, dark brown. Muscle and epithelium, pale violet. Erythrocytes and elastic fibres are stained a rich brown, while mucus, collagen are in varying shades of green ; myelin sheaths, pink ; and axis cylinders, dark blue. GIEMSA STAIN For malarial parasites, rickettsia, etc. Solution required: Giemsa stain . . . . . . i ml. Distilled water, buffered to pH 7-2 20 ml. N.B.: This mixture should be freshly prepared immediately before use. Technique: 1. Fix small pieces of tissue in 10% formalin, Regaud or Zenker. 2. Dehydrate; clear and embed. 3. Bring down paraffin sections to distilled water in the usual manner. 4. Stain for eighteen to twenty-four hours in the diluted Giemsa (as above). 5. Wash in distilled water; differentiate quickly in 0-5% acetic acid until the section is pink ; then wash with distilled water. 6. Blot and dry in air and mount. Results: Nuclei are stained dark red ; erythrocytes, pink. Malaria para- sites, bluish red with red chromatin. 105 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES GBEMSA - WRIGHT STAIN A permanent stain for differentiating the structures, particularly Nissl bodies and cytons, of the spinal cord Solution required: Wright's stain . . . . • • 5 volumes Giemsa stain . . . . . . i volume Technique: 1. Material should be fixed in neutral formalin io%. 2. Wash, dehydrate, clear, and embed in paraffin wax in the usual manner. 3. Fix sections to slides; de-wax; pass through the usual descending grades of alcohol, down to distilled water. 4. Flood the sections with a measured volume of the above staining solution and allow it to act for two minutes. 5. Add an equal volume of distilled water and mix with stain by rocking the slides gently. Allow this diluted stain to act for two minutes. 6. Pour off excess stain and immerse the slides in fresh dis- tilled water for one minute. 7. Transfer immediately into 80% alcohol and leave therein for fifteen seconds. 8. Dehydrate rapidly in 95% and absolute alcohol. 9. Clear in xylol and mount. Results: Cytons and Nissl granules are stained deep blue. Nuclei of blood-vessel structures and neuroglia are light blue. Elastic fibres of blood vessels, deep blue. Erythrocytes, pink. Neuroglia fibres, light red. Note: The proportion of the Giemsa stain regulates the inten- sity of the cyton stain. Reference: Hanburg, L. (1935), Science, 81, 364-5. I 106 SECTION TWO GOLD CHLORroE - SUBLIMATE (Cajal) For neuroglia fibres; for astrocytes in central nervous system Solutions required: A. Neutral formalin . . . . . . 15 ml. Ammonium bromide . . . . 2 gm. Distilled water . . . . . . 85 ml. B. Gold chloride (brown or yellow) 1% aqueous. C. Mercuric chloride 5% aqueous. D. Sodium hyposulphite 10% aqueous. Technique: 1 . Fresh pieces of tissue are fixed for two to twenty-one days in Solution A. 2. Frozen sections are cut 15 to 30^ thick. 3. Rinse in several changes of distilled water. 4. Immerse sections, flattened out and not lying on top of one another, for three to four hours in a freshly prepared mixture con- sisting of: Solution B . . . . • • 5 t^- Solution C . . . . • • 5 nil- Distilled water . . . . • • 30 ml. {Note: 3 ml. of the mixture is required for each section) until the astrocytes are stained dark against a relatively light background; the reaction should be controlled by microscopic examination of a section while still wet. 5. Wash in distilled water; then fix^in Solution D. 6. Wash thoroughly in tap water. 7. Dehydrate ; clear and mount. Results: Astrocytes, black. Nerve cells, red. Nerve fibres, unstained. Background, light brownish purple, or unstained. 107 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES GOLGI METHOD (Rapid) For nerve cells Solutions required: A. Potass, bichromate 3% . . . . 40 ml. Osmic acid 1% . . . . . . 10 ml. This solution must be freshly prepared. B. Silver nitrate 075%. Technique. 1. Immediately the animal is killed, tissues are cut into slices about 2 mm. thick and fixed in Solution A for one to three days depending upon the size of the pieces. 2. After removing excess fixative by blotting, the tissues are rinsed in Solution B until no more precipitate is formed. 3. Transfer to a fresh lot of Solution B and leave for two days or longer. 4. With a camel-hair brush carefully brush the precipitate from the surface of the tissue ; then wash well in distilled water. 5. Dehydrate by immersing for one to four hours, depending upon the size of the pieces, in each of the following : two lots of 95% alcohol; two lots of absolute alcohol; one lot of ether- alcohol (equal vols, ether and absolute alcohol). 6. Embed in Celloidin or L.V.N. 7. Sections 6o-ioo/^ are mounted on slides and covered with Canada balsam in benzol or cristalite, without cover glasses {see L.V.N, technique, page 26). Results: Background, dull yellow. Nerve cells and their processes, black. Blood vessels, black. GRAM'S IODINE For bacteria in sections Solutions required: A. Carbol gentian violet. B. Gram's iodine. 108 I SECTION TWO C. Carbol fuchsin (Ziehl Neelsen) . . i volume Distilled water . . . . • • 9 volumes D. Picric acid, saturated, aqueous. Technique: 1. Pieces of tissue are fixed in io% formalin; dehydrated; cleared and embedded in paraffin wax. 2. Fix sections to slides ; de-wax and take down to distilled water in the usual manner. 3. Stain in Solution A for about two minutes. 4. Pour off excess stain and without washing add Gram's iodine and allow the stain to act for one minute. 5. Differentiate in pure acetone until colour ceases to come out of the sections. 6. Counterstain in the carbol fuchsin (Solution C) for about a minute. 7. Pour off excess stain, and drain, without allowing the sections to dry; then without washing: 8. Cover the sections with the picric acid solution, pouring off after one half to one minute. 9. Dehydrate and clear with pure acetone for about fifteen seconds. 10. Clear in xylol and mount. Results: Gram-positive organisms are stained violet, while Gram- negative are red. Nuclei are stained pink, while cytoplasm is yellow. HAEMALUM - EOSIN For demonstrating collagenous tissue Solution required: A. Haemalum (Mayer). B. Eosin, yellowish 0-2% in 20% alcohol. 109 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Paraffin sections are fixed to slides, de-waxed and taken down through descending grades of alcohol to distilled water in the usual manner. 2. Stain for five to ten minutes with the haemalum solution, examining under the microscope at intervals until a satisfactory degree of staining has been achieved. 3. Rinse for a few seconds in tap water. 4. Stain for one or two seconds with the eosin solution. 5. Rinse for a few minutes in running tap water. 6. Pass through 70%, 90% and absolute, alcohol. 7. Clear in xylol and mount. Results: Collagen, deep pink. Smooth muscle, pink. Cytoplasm, pale pink. Nuclei, blue. HAEMATOXYLIN - AZOPHLOXINE For muscle, connective tissue, ganglion cells, etc. Solutions required: A. Lavdowsky's Fixative: Formalin (40% formaldehyde) . . 10 ml. Glacial acetic acid . . . . . . 2 ml. Alcohol 95% . . . . . . 50 ml. Distilled water . . . . . . 40 ml. B. Haematoxylin (Delafield, Harris or Ehrlich) C. Azophloxine . . . . . . 0*2 gm. Acetic acid 0*2% . . . . . . 100 ml. Add about i ml. chloroform as a preservative. D. Acetic acid 0-2% aqueous. E. Orange G. . . . . . . . . 2 gm. Phosphotungstic acid . . • • 4 gm. Distilled water . . . . . . 100 ml. Fast Green FCF . . . . . . 0-2 gm. no 1 SECTION TWO Technique: 1. Fix material in Lavdowsky's mixture or in io% formalin and embed in paraffin wax. 2. Stain in Harris, Ehrlich or Delafield Haematoxylin for five to ten minutes. 3. Blue in tap water or 1% lithium carbonate solution. 4. Stain in the azophloxine solution for two minutes. 5. Rinse in 0-2% acetic acid. 6. Stain in the orange G-fast green solution for one minute. 7. Differentiate for about five minutes with 0-2% acetic acid. 8. Rinse in distilled water. 9. Remove excess distilled water by draining and blotting round the edges of the sections carefully, but do not allow to dry com- pletely. 10. Dehydrate directly with two or three changes of absolute alcohol, or with cellosolve. Note: Absolute and 70% alcohol should be avoided as they have a tendency to remove the azophloxine. Results: Connective tissue, green. Striated muscle, brick red. Smooth muscle, reddish violet. Nerves, blue-grey. Ganglion cells, violet. Erythrocytes, orange. Cardiac conductive tissue is easily dis- tinguishable from cardiac muscle as it takes a lighter shade of staining. Note: Azophloxine is used here as a substitute for ponceau de xylidine in Goldner's modification of Masson's technique. The stain is also suggested in place of Eosin as a counterstain for use with haematoxylin. The advantages of using azophloxine are that it gives clear and delicate pictures and it does not overstain, if the recommended procedure is followed. When azophloxine is to be used merely as a counterstain for haematoxylin the pro- cedure is as follows : I. Proceed as steps, i, 2, 3, 4, and 5 (above). II. Rinse in distilled water. III. Dehydrate with three changes of absolute alcohol; then clear and mount. Reference: Halper, M. H. (1954, November), Stain Tech., 29, no. 6, 315-7. Ill MEDICAL AND BIOLOGICAL STAINING TECHNIQUES HAEMATOXYLIN - BASIC FUCHSIN For haemofuscin, melanin and haemosiderin in animal tissues Solutions required: A. Haematoxylin (Ehrlich). B. Basic fuchsin 0-5% in 50% alcohol. Technique: Tissues may be fixed in Zenker or in absolute alcohol or in 10% formalin. Paraffin or Celloidin sections may be employed. If Zenker's fixative is used it will be necessary to remove mercury deposits in the usual manner. 1 . Stain for five to ten minutes in Ehrlich haematoxylin. 2. Wash well in tap water, then several times in distilled water. 3. Stain from five to twenty minutes in the basic fuchsin solu- tion ; then pour off excess stain and wash well in distilled water. 4. Differentiate in 95% alcohol; then dehydrate in absolute alcohol ; clear in xylol and mount in balsam. Results: Nuclei, blue; melanin and haemosiderin remain unstained in their natural brown colours ; haemofuscin, bright red. HAEMATOXYLIN (DELAFIELD) - EOSIN For general staining Solutions required: A. Delafield haematoxylin. B. Eosin, yellowish, 1%, aqueous. Technique: I. Tissues should be fixed in Zenker, Bouin or 10% formalin and embedded in paraffin wax. 112 SECTION TWO 2. Sections are brought down to distilled water; then stained with Delafield haematoxylin for ten minutes. 3. Wash and immerse in tap water for about five minutes until the section appears blue to the naked eye. 4. Wash rapidly with distilled water ; then stain for one to two minutes with 1% eosin yellowish, aqueous. 5. Wash quickly with distilled water; then dehydrate with 95% and absolute alcohol. 6. Clear in xylol and mount. Results: Nuclei are stained blue; cytoplasm, pink. HAEMATOXYLIN (Ehrlich) For keratohyalin Solutions required: A. Haematoxylin (Ehrlich). B. Potass, permanganate o-i%. Technique: 1. Material should be fixed in 10% formalin and embedded in paraffin wax. 2. Fix sections to slides and bring down to distilled water as usual. 3. Stain in Ehrlich haematoxylin for ten minutes. 4. Pour off excess stain ; then rinse and blue in tap water, 5. Immerse in potassium permanganate o-i% for ten seconds; then wash well with water. 6. Dehydrate; clear; mount in balsam. Results: Keratohyalin is stained blue-black while the other elements are unstained or faintly stained. 113 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES HAEMATOXYLIN (Ehrlich) For demonstrating sodium urate in animal tissue Solution required: Ehrlich or Delafield haematoxylin. Technique: 1 . Fix tissues in absolute alcohol and embed in Celloidin. 2. Stain for five to ten minutes in the haematoxylin solution. 3. Blue in saturated lithium carbonate solution. 4. Rinse quickly in distilled water. 5. Dehydrate with 95% alcohol. 6. Clear in terpineol. 7. Mount in balsam. Results: Sodium urate crystals and nuclei, deep blue. HAEMATOXYLIN (Ehrlich) - EOSIN A general stain for animal tissues Solutions required: A. Haematoxylin (Ehrlich). B. Eosin, aqueous, yellowish 1% in tap water. Technique: 1. Sections are mounted on slides and brought down to distilled water in the usual manner, any mercurial deposit from the fixative being removed by the standard technique. 2. Stain from two to ten minutes in haematoxylin (Ehrlich) depending upon the " ripeness " of the stain (a well-ripened haematoxylin will act much more rapidly than a recently-prepared solution). 3. Rinse in water; then *' blue " in tap water; that is, immerse the preparation in tap water from two to ten minutes or until the preparation appears blue to the naked eye. 4. The preparation should now be examined, while still wet, 114 \ SECTION TWO under the microscope and if the nuclei are not stained a bright and transparent blue and the cytoplasm (except mucin and basophile granules of mast cells, etc.) is not colourless, rinse in water and repeat the staining and bluing process. If, on the other hand, overstaining has taken place, immerse the preparation in 0-5% HCl for five to thirty seconds; then immedi- ately rinse in water, " blue " in tap water and again examine under the microscope; if the sections are still overstained repeat the treatment with HCl; rinse and " blue " in tap water again. 5. Wash well with water; then stain from two to five minutes in eosin solution. 6. Rinse quickly in water and examine the section rapidly, while still wet, under the microscope to ensure that the depth of the counterstain is sufficient. The cell cytoplasm, collagen, connec- tive tissue fibres, erythrocytes, etc., should be stained a bright transparent pink. It is advantageous to overstain somewhat with the eosin, as subsequent dehydration in the alcohols will remove the excess eosin. 7. Dehydrate by passing rapidly through 70%, 90% and abso- lute alcohol. 8. Clear in xylol ; mount in balsam. Results: Nuclei are stained dark blue; karosomes, dark blue; plasma- somes, red ; cytoplasm (except when basophile) is stained in varying shades of pink; muscle and collagen fibres, pink; elastic fibres (when thick), pink; erythrocytes, thyroid colloid and keratin, bright pink. HAEMATOXYLIN - FLUORCHROME (Kultschitzky-Pal) For myelin sheaths. Particularly suitable for demonstrat- ing very fine fibres in cerebal cortex, etc. Solutions required: A. Weigerfs rapid fixative : Fluorchrome . . . . . . 2 gm. Potassium bichromate . . • • 5 gni- Distilled water . . . . . . 100 ml. Add the potassium bichromate to the water ; boil ; add the fluorchrome; cool; then filter. 115 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES B. Kultschitzky's haematoxylin solution. C. Lithium carbonate, saturated, aqueous . . . . . . lOO ml. Potassium ferricyanide i% aqueous . . . . . . lo ml. Technique: 1 . Slices of formalin-fixed tissue not more than 5 mm. thick are immersed in Weigert's rapid fixative from four to seven days. 2. Wash in running water for four to six hours. 3. Dehydrate and embed in Celloidin. 4. Stain sections in Kultschitzky's haematoxylin from twelve to twenty-four hours. 5. Differentiate for several hours in Solution C controlling under the microscope at intervals of one half to one hour, until the white matter is stained blue-black, and the grey matter is stained yellowish. Note: For human spinal cord differentiation takes up to twelve hours. Human cerebral cortex requires about four hours. 6. Wash well in running water. 7. Dehydrate and clear by the standard Celloidin method. 8. Mount in cristalite. Results: Myelin sheaths are stained black, while ground-substance is yellow. HAEMATOXYLIN - GENTIAN VIOLET - IODINE For demonstrating Gram-positive bacteria and fibrin in sections Solutions required: A. Haematoxylin (Delafield). B. Aniline gentian violet C. Lugol's iodine. D. Aniline oil. . .. .. .. 20 ml. Xylol . . . . . . . . 10 ml. E. Erythrosin 5% in absolute alcohol. 116 SECTION TWO Technique: 1. Tissues may be fixed in io% formalin or in Zenker, if the latter is used then mercuric precipitates must be removed from the sections by the standard technique. 2. Fix sections to slides ; de-wax with xylol and pass through descending grades of alcohol to water in usual manner. 3. Stain in the haematoxylin solution for five to twenty minutes. 4. Rinse quickly in acid alcohol. 5. Immerse in a large volume of tap water for two to five min- utes. 6. Stain in the aniline gentian violet for two to five minutes. 7. Pour off excess stain, and without washing, blot the slide carefully. 8. Flood with Lugol's iodine and allow the solution to act for two to five minutes. 9. Pour off excess, and without washing, blot dry carefully. 10. Decolorize for a few seconds with Solution D. 11. Flood the preparation with erythrosin (Solution E, above) and allow the stain to act from one half to one minute. 12. Pour oflF excess stain and wash the preparation with Solution D. 13. Rinse well with xylol; drain off excess and blot dry care- fully. 14. Mount in balsam or Cristalite or Clearmount. Results: Nuclei, blue. Fibrin and Gram-positive organisms, purplish blue. HAEMATOXYLIN (Heidenhain) Solutions required: A. Iron alum 3 gm. Distilled water . . . . . . 100 ml Dissolve by shaking. 117 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES B. Haematoxylin io% in absolute alcohol (which has been ripened for three months or longer) . . 5 ml. Distilled water . . . . • • 95 r^l- C. Eosin yellowish 1% aqueous. Technique: Fix in Zenker. Embed in paraffin wax. 1. Sections are brought down to distilled water, then mor- danted in Solution A for one half to three hours. 2. Wash in tap water ; stain one to three hours in Solution B ; then rinse in tap water. 3. Differentiate in Solution A, controlling by examination under the microscope. 4. Wash in running water for five to ten minutes. 5. Stain with Solution C for one to three minutes. 6. Wash in tap water; dehydrate; then clear in xylol and mount. Results: Nuclei are stained black; cytoplasmic structures, pink. HAEMATOXYLIN For the identification of lipines Solutions required: A. Potassium bichromate 5% aqueous. B. Haematoxylin solution (Ehrlich). C. Potassium ferricyanide . . . . 2-5 gm. Borax . . . . . . . . 2 gm. Distilled water . . . . . . 100 ml. Technique: I. Tissues are fixed from twelve to twenty-four hours in 10% formalin in normal saline. 118 SECTION TWO 2. Wash for several hours in running water. 3. Make frozen sections and collect them in distilled water. 4. Immerse in Solution A for twenty-four to forty-eight hours at 37° C. 5. Wash in several changes of distilled water, handling the sec- tions with care (as they become brittle after immersion in Solution A). 6. Immerse in Solution B for four to six hours at 37° C. 7. Wash in distilled water. 8. Differentiate in Solution C, controlling under the micro- scope, until the ground cytoplasm is changed from black to yellow. This process takes several hours. 9. Wash thoroughly in five or six changes of distilled water; then mount in glycerine jelly. Result: Lecithin and other lipines are stained black to deep blue (light blue coloration should not be taken as positive). Lipides and other tissue constituents are colourless. HAEMATOXYLIN (Kultschitzky) (Weigert's modification) For finer studies of cortical architecture and for total brain sections Solutions required: A. Weigerfs Secondary Mordant: Cupric acetate neutral, normal . . 5 gm. Fluorchrome . . . . . . 2-5 gm. Distilled water . . . . . . 100 ml. Boil ; allow to cool ; then add : Glacial acetic acid . . . . 5 ml. B. Haematoxylin (Kultschitzky). C. Lithium carbonate, saturated, aqueous . . . . . . 100 ml. Potassium ferricyanide 1% aqueous 10 ml. K 119 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. After fixing material in io% formalin mordant for four to hwt days in Solution A. 2. Dehydrate in ascending grades of alcohol in the usual way, and embed in Celloidin. 3. Immerse for twelve to twenty-four hours in the haematoxylin (Solution B). 4. Differentiate in Solution C from four to twelve hours, con- trolling by examination under the microscope at intervals, and changing the differentiating fluid three or four times. 5. Wash thoroughly in distilled water. 6. Dehydrate with 95% alcohol. 7. Clear in terpineol. 8. Drain well and blot carefully. 9. Mount in balsam. Results: Finest myelin sheaths are stained a deep black. HAEMATOXYLIN - PHLOXINE - ANILINE GENTIAN VIOLET For actinomyces in sections Solutions required: A. Ehrlich haematoxylin. B. Phloxine 3% aqueous. C. Aniline gentian violet. D. Gram's iodine. Technique: 1. Tissues are fixed in 10% formalin, washed, dehydrated, cleared and embedded in paraffin wax in the usual manner. 2. Fix sections to slides ; de-wax and bring down to distilled water as usual. 3. Stain with Ehrlich haematoxylin five to ten minutes; then blue and wash in lithium carbonate saturated aqueous. 120 SECTION TWO 4. Stain for fifteen to twenty-five minutes in the phloxine solution; then wash with distilled water. 5. Stain in aniline gentian violet for about ten minutes. 6. Rinse in distilled water. 7. Immerse in Gram's iodine solution for one minute. 8. Wash in distilled water. 9. Decolorize with aniline oil until the stain ceases to come out of the sections. 10. Rinse well in several changes of xylol and mount. Results: Branched forms are stained blue, while clubs appear red. HAEMATOXYLIN, PHOSPHOTUNGSTIC (Mallory) For Pleuropneumonia organisms in sections of lung Solution required: Phosphotungstic acid haematoxylin (Mallory). Technique: 1. Pieces of tissue should be fixed in Bouin, Carnoy or absolute alcohol and embedded in paraffin wax in the usual manner. 2. Fix sections to slides ; de-wax, pass through descending grades of alcohol down to distilled water, as usual. 3. Stain in Mallory's phosphotungstic acid haematoxylin in a stoppered staining jar for twenty-four hours, without treatment with the usual potassium permanganate and oxalic acid solutions. 4. Pour off excess stain ; drain well without washing ; then care- fully blot dry. 5. Dehydrate quickly in absolute alcohol. 6. Clear in xylol and mount. Results: The organisms appear as masses of mycelia which are stained deep blue. 121 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES HAEMATOXYLIN - PICRO FUCHSIN For nuclei, connective tissue, etc. Solutions required: A. Distilled water . . . . . . 47-5 ml. Ferric chloride, hydrated 4% aqueous . . . . . . . . 2 ml. Hamaetoxylin 10% in absolute alco- hol . . . . . . . . . . 0*4 ml. B. Picric acid, saturated, aqueous . . 20 ml. Acid fuchsin 1% aqueous . . . . 0-5 ml. C. Picric acid, saturated in absolute alcohol. Technique: 1 . Tissues are fixed in Bouin and embedded in paraffin wax. 2. Sections about 8ju, in thickness are fixed to slides, dewaxed with xylol and taken through the usual descending grades of alcohol to distilled water. 3. Stain for two to three minutes in solution A. 4. Differentiate and counterstain for about ten to fifteen seconds in solution B, controlling under the microscope, until only the nuclei are stained a greyish colour with the haematoxylin. 5. Rinse immediately in distilled water. 6. Dehydrate by dripping solution C onto the slide. 7. Clear with Terpineal. 8. Mount directly with Michrome mountant, or rinse with xylol, then mount with Clearmount or Cristalite. Results: Chromatin, black to grey. Muscle, yellow. Connective tissue, red. Keratinized regions, bright yellow. Cytoplasm, yellow. Reference: Margolena, L. A. and Dolnick, E. H. (1951), Stain Tech., vol. 26, pp. 1 1 9-2 1. 122 SECTION TWO HAEMATOXYLIN - PICRO PONCEAU S A selective stain for collagen and connective tissue in place of Haematoxylin - Van Gieson Solutions required: A. Haematoxylin (Heidenhain or Ehrlich). B. Picro Ponceau S {Curtis): Ponceau S i% aqueous . . . . lo ml. Picric acid i% aqueous . . . . 86 ml. Acetic acid i% aqueous . . . . 4 ml. Technique: Proceed exactly as for Haematoxylin (Ehrlich) - Van Gieson or Haematoxylin (Heidenhain) - Van Gieson (pages 127-128). Results: Identical with Haematoxylin - Van Gieson. Note: Unlike Van Gieson, Picro Ponceau does not fade when mounted in Canada balsam: but Van Gieson does not fade when mounted in D.P.X. or Clearmount or Cristalite. HAEMATOXYLIN (Weigert) - PONCEAU FUCHSIN (Curtis) Note: The chief advantages of this method over numerous other trichrome techniques, is its simplicity and reliability and the fact that it works well after any fixative. The disadvantage is that cytoplasmic details are not as clearly revealed as after such stains as Masson or Mallory. Solutions required: A. Weigert's Haematoxylin, A B. Weigert's Haematoxylin, B C. Curtis Stain: Ponceau S, 2% aqueous . . • • 5 ml. Picric acid, saturated, aqueous . . 95 ml. Acetic acid 2% . . . . . . 2 ml. Technique : 1 . Fix material with any desired fixative, embed and section as usual. 2. Fix sections to slides and take down to 70% alcohol as usual. 3. If the fixative contains a salt of mercury, remove mercurial precipitate in the usual way. 123 • MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 4. Wash with distilled water. 5. Stain for five to ten minutes in a mixture consisting of one volume of each of solutions A and B, and two volumes distilled water. 6. Wash for five minutes in running water. 7. Stain for two to four minutes in Curtis Stain. 8. Wash with 95% alcohol. 9. Dehydrate with absolute alcohol. 10. Clear in xylol and mount. Results: Chromatin, black. Cytoplasm, yellow. Collagen and fibres, red. Reference: Leach, E. H. (1946), Stain Tech., 21, 107-10. HAEMATOXYLESr - PONCEAU FUCHSIN - FAST GREEN FCF A micro-anatomical stain, superior to haematoxylin and eosin for the differentiation of tissues in histo-pathological work, superior to Van Gieson for collagen fibres. The stain has also been found very valuable in cytological work Solutions required: A. Iron haematoxylin (Heidenhain) No. i. B. Iron haematoxylin (Heidenhain) No. 2. C. Picric acid 8% in 96% alcohol. D. Acid fuchsin 1% aqueous . . 100 ml. Glacial acetic acid . . . . i ml. E. Ponceau de xylidine . . . . i gm. Distilled water . . . . . . 100 ml. Glacial acetic acid . . . . i ml. F. Phosphomolybdic acid 1% aqueous. G. Fast Green FCF 2% aqueous . . 100 ml. Glacial acetic acid . . . . 2 ml. Technique: I. Bouin, formalin or Susa-fixed tissues are embedded and sectioned in the usual manner. m 124 SECTION TWO 2. Immerse sections, which have been brought down to distilled water as usual, for one half to one hour in iron haematoxylin (Heidenhain) No. i. 3. Rinse well in distilled water. 4. Stain in iron haematoxylin (Heidenhain) No. 2 for a length of time equal to the duration of stage 2 (above) ; then rinse thoroughly in distilled water. 5. Differentiate in solution C until only the nuclei are stained; then wash well in water ; and stain in solution D for five minutes : wash in distilled water. 6. Stain one to five minutes in a mixture consisting of one part of Solution E (above) and nine parts of distilled water ; then rinse thoroughly in tap water. 7. Differentiate in 1% phosphomolybdic acid for five to fifteen minutes until collagen fibres are almost colourless; then, without rinsing : 8. Stain in the Fast Green FCF solution for half to two min- utes. 9. Differentiate by washing in water. 10. Dehydrate in the usual manner; clear and mount. Results: Nuclei are stained mauve to black; cytoplasm, varying shades of red and mauve ; muscle, red ; collagen fibres and mucus, green. HAEMATOXYLIN - PONCEAU S - PICRO ANILINE BLUE For differential staining of connective tissue and muscle Solutions required: A. Haematoxylin (Weigert) A. B. Haematoxylin (Weigert) B. C. Ponceau, S 0-2% aqueous . . 100 ml. Glacial acetic acid . . . . i ml. D. Picric acid, saturated aqueous . . 100 ml. Aniline blue, water soluble . . o-i gm. E. Acetic acid 1% aqueous. 125 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Tissues should be fixed in io% formalin and paraffin sections employed. 2. Stain for five minutes in a freshly prepared mixture consisting of equal parts of Weigert's Haematoxylin A and B. 3. Wash in tap water. 4. Stain for three to five minutes in the acetic Ponceau, S (Solution C). 5. Rinse in distilled water. 6. Stain for three to five minutes in the picro aniline blue (Solution D). 7. Wash for three or four minutes in 1% acetic acid solution. 8. Dehydrate in ascending strengths of alcohol and clear in xylol in the usual manner. 9. Mount in acid balsam. Results: Connective tissue, glomerular basement membrane and reti- culum, blue. Muscle and plasma, pink. Erythrocytes, bright red. HAEMATOXYLIN (Weigert) SCARLET R For demonstrating fatty acids crystals, soaps and neutral fats in fat necrosis Solutions required: A. Formalin 10% saturated with calcium salicylate. B. Copper acetate 10% aqueous. C. Weigert haematoxylin, A. D. Weigert haematoxylin, B. E. Borax 0-2% aqueous . . . . i litre Potassium ferricyanide . . . . 2-5 gm. F. Scarlet R (Herzheimer). Acetone . . . . . . • • 50 ml. Alcohol, 70% . . . . . . 50 ml. Scarlet R . . . . . . . . i -5 gm. Heat on a hot water bath; then allow to cool before filtering. 126 SECTION TWO Technique: 1. Fix tissues in Solution A; wash in running water and cut frozen sections. 2. Mordant the sections in Solution B for three to twenty-four hours; then wash in water. 3. Immerse in mixture of equal parts of Solutions C and D for twenty to forty-five minutes. 4. Differentiate in Solution E, examining under the microscope at intervals. 5. Wash well with distilled water. 6. Stain with Solution F for about five minutes. 7. Rinse quickly with 70% alcohol. 8. Rinse with distilled water. 9. Mount in neutral glycerine jelly. Results: Neutral fats are stained red, whilst fatty acids are deep blue black and haemoglobin, calcium and iron may also be stained. Note: Calcium salicylate is added to the formalin fixative to convert soaps, which are sodium and potassium salts of fatty acids, into insoluble calcium soaps. If it is desired to demonstrate how much, if any, soap is present in addition to fatty acids, com- pare stained sections of two pieces of the same material, fixing one piece in Solution A and the other in ordinary 10% formalin. HAEMATOXYLIN (Ehrlich) - VAN GIESON STAIN A selective stain, for collagen and connective tissue, w^hich requires less time than the Haematoxylin (Heidenhain) - Van Gieson technique, although the results are not as satisfactory Solutions required: A. Haematoxylin (Ehrlich). B. Van Gieson stain (Picro Acid fuchsin). Technique : I . Mount sections on slides ; dewax and pass through the usual descending grades of alcohol to water. 127 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 2. Stain in Ehrlich haematoxylin for ten to thirty minutes. 3. Rinse in water. 4. Blue in tap water or 1% lithium carbonate solution. 5. Stain for three to five minutes in Van Gieson stain. 6. Rinse for one or two seconds in water. 7. Drain and draw off excess water by applying a piece of filter paper to the edges of the section. 8. Dehydrate with absolute alcohol only. 9. Clear in xylol and mount in D.P.X., or Cristalite or Clear- mount. Note: (a) It is essential, at stage 2, to overstain with haematoxylin as this stain is differentiated by the picric acid of the Van Gieson stain. (b) Preparations mounted in Canada balsam fade, but fading can be obviated by use of one of the recommended mountants. HAEMATOXYLIN (Heidenhain) - VAN GIESON STAIN A selective stain for collagen and connective tissue, superior to Haematoxylin (Ehrlich) - Van Gieson Solutions required: A. Haematoxylin (Heidenhain) A. B. Haematoxylin (Heidenhain) B. C. Van Gieson stain. Technique: 1. Fix pieces of tissue in Bouin, Carnoy, Susa or 10% formalin. 2. Fix sections to slides; dewax and take down to water in the usual way, after removing mercurial precipitate if Susa has been used as the fixative. 128 SECTION TWO 3. Immerse in solution A for one half to one hour. Note : If a fixative other than Bouin, Carnoy, Susa or formaUn has been used it will be necessary to increase the time in solution A and in solution B up to twelve hours or longer : the time varies for different fixatives. 4. Rinse in water. 5. Stain in solution B for a time exactly equal to step 3. 6. Rinse in water. 7. Differentiate with solution A, controlling by examination under the microscope, after the preparation has been rinsed briefly in water. 8. Wash gently in running water for about five minutes to remove all traces of solution A (iron alum). 9. Stain for three to five minutes in Van Gieson. 10. Rinse for a few seconds in water. 11. Examine, while still wet, under the microscope. 12. Continue the staining with Van Gieson, or continue the differentiation with water, whichever is necessary. 13. Drain and draw off excess water by means of a filter paper applied carefully to the edges of the section, but do not allow the preparation to dry completely. 14. Dehydrate with absolute alcohol only. 15. Clear in xylol. 16. Mount in D.P.X., Cristalite or Clearmount. Results: Nuclei of cells: dark brown to black. Collagen fibres: bright red. Erythrocytes, muscle, epithelia and other tissues: yellow. Note: Van Gieson stain fades if mounted in Canada balsam, but fading can be avoided by the use of D.P.X., or Cristalite or Clear- mount. 129 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES mCKSON'S PURPLE A general stain suitable for class work Solution required: Hickson purple, saturated aqueous. Technique: 1 . Bring sections down to water as usual. 2. Stain in Hickson's purple for ten to twenty minutes. 3. Dehydrate, clear and mount. Results: Leucocytes, purple ; erythrocytes, distinct red. The rest of the tissues purple. Reference: Cannon, H. G. (1941),^- Roy. Micr. Soc, series III, 61, parts 3 and 4. HICKSON'S PURPLE - VICTORIA GREEN A general stain, particularly suitable for class work Solutions required: A. Hickson's Purple saturated aqueous. B. Victoria green, G. saturated in 70% alcohol. Technique: 1 . Fix sections to slides ; dewax and take through the alcohols down to distilled water as usual. 2. Stain for ten minutes in the Hickson's purple. 3. Rinse in distilled water. 4. Stain in the victoria green for half to one hour. 130 SECTION TWO 5. Rinse in 70%, followed by 90% alcohol. 6. Clear in xylol and mount. Results: Nuclear are sharply defined, purple. Erythrocytes sharply defined, stained vivid green, against a general blue-purple back- ground. Note: This method is an improvement of Hickson purple used alone. Reference: Cannon, H. G. (1941), J. Roy. Micr. Soc, series III, 61, parts 3 and 4. fflTCHCOCK AND EHRLICH'S MIXTURE For lymphatic, ganglion, plasma and basophilic cells; im- mature cells of bone marrow, striated muscle, and fibrin. Technique: 1. Fix in Zenker-acetic acid, corrosive sublimate, but not in Miiller or formalin. 2. Paraffin sections are brought down to 90% alcohol; then passed through a solution of iodine in 90% alcohol. 3. The iodine is removed by passing through the graded alcohols to water and finally washing for fifteen minutes in running water. 4. Flood with the stain and allow it to act for fifteen to thirty seconds ; then pour oif the stain and wash rapidly in water. 5. The preparation is then passed directly into absolute alcohol, where it is allowed to remain only as long as the stain continues to be washed out in clouds. 6. Clear in xylol and mount. Note: Sometimes the brilliancy of the stain is enhanced by re-staining. Results: Plasma cells: cytoplasm, brilliant crimson: nuclei, bluish- green. Other cells appear in lighter shades of green and crimson. 131 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES JENNER STAIN For blood-forming organs Solutions required: A. Formol- Saline. Formalin, cone. (i.e. 40% for- maldehyde) 100 ml. Sodium chloride, A.R. . . 8-5 gm. Distilled water I litre Acid sodium phosphate, mono- hydrate, A.R. . . 4 giAi. Anhydrous disodium phosphate. A R >£x»XX* •• •• •• •• 6-5 gm. B. Jenner stain. Technique: 1 . Fix pieces of tissue for two or three days in Solution A. 2. Dehydrate in ascending grades of alcohol as usual; clear; embed in paraffin wax. 3. Fix sections, not exceeding 5/z in thickness, to slides ; de-wax ; pass through the usual descending grades of alcohol down to dis- tilled water which has been buffered to pH 7-0. 4. Stain for forty-five minutes in a grooved, stoppered staining jar, with a mixture consisting of equal volumes of Jenner stain and distilled water, buffered to pH 7.0. 5. Differentiate and dehydrate with absolute alcohol. 6. Clear in xylol and mount in Cristalite. Results: Neutrophile granules are stained pink. Oxyphile granules, brownish red. Basophile granules, purple. Nucleoli (plasmosomes), pink. The cytoplasm of partially haemoglobinated precursors of erythrocytes are stained in varying shades of reddish violet, while mature erythrocytes are deep pinkish orange. 132 SECTION TWO JENNER STAIN - GIEMSA STAIN For the polychromatic staining of blood-forming organs Solutions required: A. Formol saline. B. Jenner Stain. C. Giemsa stain . . . . . . i ml. Distilled water (buffered to pH 70) 20 ml. D. Acetic acid 008% aqueous Technique: 1 . Fix material in Solution A (above) from twelve to forty-eight hours. 2. Dehydrate in the alcohols and clear as usual; embed in paraffin wax and cut sections not exceeding 5^ in thickness. 3. Fix sections to slides and remove wax with xylol. 4. Wash well with two changes of pure methyl alcohol. 5. Stain sections with a measured volume of Jenner stain, which should be freshly filtered. 6. Cover the slides with a Petri dish lid lined with two or three sheets of moistened filter paper (this is to prevent the evapor- ation of the alcohol and the consequent formation of a precipitate on the sections), and allow the stain to act for three minutes. 7. Add a volunle of distilled water (buffered to pH 7-0), equal to that of the stain, to the slides, which should now be gently rocked to ensure thorough mixing of the stain and water. 8. Allow this diluted stain to act for one minute. 9. Pour off excess stain; then without washing, immerse the slides in a stoppered staining jar containing diluted Giemsa stain (Solution C above) and leave the stain to act for forty-five minutes. 10. Rinse and differentiate in Solution D. 1 1 . Rinse thoroughly in distilled water. 12. Dehydrate quickly in 95% alcohol, followed by two changes of absolute alcohol. 133 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 13. Clear in xylol and mount in Cristalite. Results: Erythroc3rtes are stained orange. Cytoplasm of lymphocytes and blastocytes are blue. Nuclei, deep blue to violet. Mast cell granules, violet to violet-red. LEAD HAEMATOXYLIN - ACID FUCHSIN (MacConaill) A " definitive '* polychrome stain for the central nervous system and the trunks outside it. The essence of this technique, which is due to Professor M. A. MacConaill, of the Department of Anatomy, University College, Cork, Ireland, is that the lead haematoxylin reduces the ammon- ium molybdate to form a blue lake, which makes it possible to employ only the minimum exposure to haematoxylin, thereby leaving the erythrophile (*' Fuchsinophile ") parts of the neurone red. Solutions required: A. Lead nitrate Glacial acetic acid Acid fuchsin Water B. Haematoxylin Acetic acid 4% aqueous C. Solution A Solution B 2gm. 8 ml. 0-5 gm. 92 ml. I gm. 100 ml. I volume I volume Note: This solution should be prepared as and when required : it deteriorates after one day. D. Liquor ammon. acetat. B.P. . . 10 ml. Ammonium molybdate, saturated, aqueous.. .. .. .. 70 ml. Water . . . . . . . . 80 ml. Note: All the above solutions must be made with- out the application of heat. Tap water may be used : the solutions must be filtered. 134 SECTION TWO Technique: 1. Material should be fixed in 5 or 10% formalin and embedded in paraffin wax. Sections are cut 6 to 12^ in thickness. 2. Fix sections to slides; remove paraffin wax and take down to 70% alcohol by the usual stages. 3. Pass through 30% alcohol; then stain in Solution C for five minutes. 4. Rinse in two changes of tap water. 5. Immerse in Solution D for one to two minutes. 6. Wash in running water for two to five minutes to remove the unchanged molybdate. 7. Dehydrate; clear in xylol and mount. Note: A deep yellow filter is of great help in microscopic examination, although not necessary. Results: Nuclei, dark blue ; nucleoli of neurones, red ; axial substances of nerve fibres, dark to pale blue ; cuticular substance (including myelotheca) of nerve fibres, red ; neurilemma (of Glees), purplish red. Note: To eliminate all myelin, sections should be passed through Cellosolve after the alcohols. The same precaution should be observed when preparing tissue for embedding. From Proceedings of the Royal Irish Academy, Vol. 53, Section B, No. i, The Myelothecal Apparatus of Human Nerve; and from personal communications with Professor M. A. MacConaill, m.r.i.a. LEISHMAN STAIN For general differentiation of blood corpuscles; for malarial parasites; trypanosomes, etc. This stain is extensively used by British workers who generally prefer it to Wright's stain which is used extensively in America. Solutions required: A. Formol saline, neutral, buflFered. B. Leishman stain. C. Acetic acid o-o8% aqueous. L 135 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1 . Fix pieces of tissue in Solution A for sixteen to forty-eight hours. 2. Dehydrate in the usual ascending grades of alcohol ; clear ; and embed in paraffin wax. 3. Fix sections, not exceeding 5^ in thickness to slides; remove wax with xylol ; pass through descending grades of alcohol down to neutral distilled water. 4. Stain for five to ten minutes in freshly prepared mixture consisting of one volume of Wright's stain and two volumes of neutral distilled water, in a stoppered staining jar. 5. Rinse with neutral distilled water. 6. Differentiate with the acetic acid solution, controlling by examination under the microscope, until the protoplasm of the cells is pink, and only nuclei are blue. 7. Wash with neutral distilled water. 8. Dehydrate quickly with absolute alcohol; clear in xylol; mount in Cristalite. Results: Erythrocytes, yellowish red. Polymorphonuclears : dark purple nuclei, reddish violet granules, pale pink cytoplasm. Eosinophiles : blue nuclei, red to orange-red granules, blue cytoplasm. Baso- philes : purple to dark blue nuclei, dark purple to black granules. Lymphocytes : dark purple nuclei, sky blue cytoplasm. Platelets : violet to purple granules. Malarial parasites and Leishmanial chromatin, red ; cytoplasm, blue. Trypanosomes : chromatin, red. Note: The timing of the staining either before or after dilution may be altered to suit individual requirements. Staining effects similar to Giemsa are obtained by staining for ten minutes in Leishman stain diluted with twice its volume of distilled water buffered to pH 6-5. LEUCO PATENT BLUE For the identification of haemoglobin. Solutiofis required: A. Patent blue AF54 (Michrome) . . i gm. Distilled water . . . . . . 100 ml. 136 SECTION TWO Dissolve; then add: Zinc metal powder . . . . lo gm. Glacial acetic acid . . . . 2 ml. Boil until the blue colour completely disappears. Allow to cool ; shake with about i gm. of decolorizing carbon; then filter. The liquid which should then be quite colourless is stored in a well stoppered bottle. B. Solution B . . . . . . 10 ml. Glacial acetic acid . . . . 2 ml. Hydrogen peroxide 3% . . i ml. N.B. : This solution must be freshly prepared and filtered before use. C. Safranin o-i% aqueous . . 99 ml. Glacial acetic acid . . . . i ml. Technique: 1. Fix tissue blocks, not more than 3 to 5 mm. in thickness in 10% formalin buffered to pH 7-0 for 24 to 28 hours (prolonged fixation should be avoided). 2. Embed in paraffin wax as usual and cut section 5 to 6/x in thickness. 3. Take sections down to water as usual. 4. Stain in solution B for three to five minutes. 5. Wash briefly in water. 6. Counterstain for thirty to sixty sections in the safranin solu- tion. 7. Rinse briefly with water. 8. Dehydrate as usual. 9. Clear in xylol. 10. Mount in Clarite or other synthetic mountant such as D.P.X., Clearmount, etc. Results: Haemoglobin, dark blue-green ; background light pink. Note: Blood and tissue smears fixed with methyl alcohol may also be stained by applying the stains as prescribed above. Reference: Adapted from Dunn, R. C. (1946), Stain. Tech., 21, 65. MEDICAL AND BIOLOGICAL STAINING TECHNIQUES LEVADITI'S STAIN For Treponema pallidum in sections Solutions required: A. Silver nitrate . . . . . . 2-5% aqueous B. Reducing solution: Pyrogallic acid . . . . • • 3 grn. Formalin . . . . . . • • 5 ml. Distilled water . . . . . . 100 ml. Technique: Tissues about i mm. thick should be fixed for twenty-four hours in 10% formalin and embedded in paraffin wax after staining. 1. After rinsing tissues in tap water, immerse in 95% alcohol for twenty-four hours. 2. Immerse in distilled water until the tissue sinks to the bottom of the jar. 3. Transfer to Solution A for 3 to 6 days at 37° C. in the dark, changing the solution every twenty-four hours. 4. Wash in distilled water; then immerse in Solution B for twenty-four to seventy-two hours in the darkroom at room tem- perature. 5. Wash in distilled water; then dehydrate with 80%, 95%, and absolute, alcohol. 6. Clear in cedarwood oil and embed in paraffin wax. 7. Sections are cut 5/^ in thickness and mounted after removal of the paraffin wax. Results: Treponema, jet black. Tissue, yellow to brown. LIGHT GREEN - ACID FUCHSIN (Alzheimer) For demonstrating neuroglia changes Solutions required: A. Osmic acid 2% aqueous . . 20 ml. Chromic acid 1% aqueous . . 75 ml. Glacial acetic acid . . . . 0-5 ml. 138 SECTION TWO B. Acid fuchsin 25% aqueous. C. Picric acid, saturated, alcoholic 15 ml. Distilled water . . . . . . 30 ml. D. Light green 10% aqueous. Technique: 1. Fix thin slices of the material in 10% formalin for twenty- four hours to three days. 2. Wash for twenty-four hours in running water. 3. Immerse very thin slices of the material in a comparatively large volume of Solution A which should be changed once or twice if it blackens. 4. Wash for several hours in running water. 5. Pass through ascending grades of alcohol. 6. Clear in the usual manner and embed in paraffin wax. 7. Sections not more than 2 to 4/^ in thickness are fixed to slides. 8. De-wax with xylol. 9. Rinse thoroughly with absolute alcohol and pass through the usual descending grades of alcohol down to distilled water. 10. Stain for an hour at 60° C. with the acid fuchsin solution. 11. Allow the preparation to cool to room temperature; then wash with water. 12. Immerse in Solution C (picric acid) from one second to two minutes. 13. Rinse in two changes of water. 14. Stain from one half to one hour in the Light Green solution. 15. Rinse quickly in absolute alcohol. 16. Rinse in xylol. 17. Mount in Canada balsam or Cristalite. Results: Migrating astrocytes, of varying shades of green and sometimes containing fuchsinophile granules of brown stained lipoid inclu- sions. Lipoid contents of perivascular phagocytes are brown to 139 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES black. Neuroglia fibres and erythrocytes, red. Medullary sheaths are unstained. Connective tissue, deep green. Nerve cells are pale green with red stippling, while nerve-cell nuclei are a darker green with bright red nuclei. Notes: (a) The material must be fresh and only small pieces should be employed. {b) Sections stained by this technique should appear lilac in colour to the naked eye. {c) It is advantageous to experiment in order to determine the optimum staining time in the picric acid and the light green, as results vary according to the material to be stained. From Pathological Technique by F. B. Mallory, by courtesy of Messrs. W. B. Saunders Co., Philadelphia, U.S.A LIGNIN PINK For whole mounts of marine invertebrates, particularly for crustaceans limbs, ostracod appendages, Medusa of Obelia, etc., as well as for demonstrating chitin Overstaining with lignin pink is impossible, and it will not wash out with alcohol. Solutions required: A. Sea water Bouin: Sea water saturated with picric acid 75 ml. Formaldehyde 40% . . . . • • 25 ml. Glacial acetic acid . . . . • • 5 ^' B. Lignin pink saturated in distilled water or in Benzyl alcohol. Technique: 1. Specimens are fixed from eighteen to forty-eight hours, according to the material, in Solution A. 2. Wash out the fixative with 50% alcohol, followed by 70% alcohol until the yellow coloration, due to the picric acid, is com- pletely extracted. 3. Wash in running water to remove the alcohol. 4. Immerse in solution B for fifteen minutes or longer. 140 SECTION TWO Results: With the aqueous solution of the stain Medusa of Obelia and limbs of crustaceans are stained deep carmine colour. The finest structures of ostracod appendages, uniform pink, but a better effect can, however, be obtained by staining the specimen for a longer period (up to sixteen hours) with a solution of the dye in benzyl alcohol : the final result in this case is a definite purple for the exoskeleton, while the other tissues are carmine colour. Reference: Cannon, H. G. (1941),^. R. Mic. Soc, series III, 61, parts 3 and 4. LITHIUM SILVER (Laidlaw) For staining skin and tumours Solutions required: A. Iodine 1% in absolute alcohol. B. Sodium thiosulphate 5% aqueous. C. Potassium permanganate 0-5% aqueous. D. Oxalic acid 5% aqueous. E. Lithium silver: Dissolve 12 gm. silver nitrate in 20 ml. distilled water in a 500 ml. stoppered bottle ; then add 230 ml. lithium carbonate, saturated, aqueous, and shake well. Transfer to a 250 ml. measuring cylinder; cover with a watch glass and allow to stand undisturbed until the precipitate formed measures about 70 ml. Pour off the clear liquid and transfer the precipitate to another vessel. Wash precipitate with three or four changes of distilled water, decanting after each washing so that the precipitate remaining measures 70 ml. Add a diluted ammonia solution (15 ml. strong ammonia solution, sp. gr. 0-880 diluted with 35 ml. distilled water) a little at a time until the fluid precipitate is almost clear. Filter through a Whatman No. 40 filter paper. F. Formalin 1% in tap water. G. Gold chloride (yellow) 0-5% in distilled water. Technique: 1. Fix tissues in 10% formalin for three days. 2. Dehydrate, clear, embed in paraffin wax in the usual manner. 141 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 3. Fix sections to slides, de-wax and take down to water as usual. 4. Wash in running water for five minutes. 5. Immerse in the iodine solution for three minutes. 6. Pour off excess iodine and immerse in the sodium thio- sulphate solution for three minutes. 7. Rinse in tap water ; then immerse in the potassium perman- ganate solution for three minutes. 8. Rinse in tap water. 9. Immerse in the oxalic acid solution for five minutes. 10. Wash in running tap water for ten minutes. 1 1 . Immerse in three changes of distilled water for three or four minutes in each. 12. Stain in an oven for five minutes with the lithium silver solution heated to 50° C. 13. Rinse the slide back and front with distilled water to remove all traces of excess lithium silver. 14. Immerse slide in ajar of 1% formalin. 15. Rinse both sides of the slide with distilled water to remove all traces of the formalin solution. 16. Immerse in the yellow gold chloride solution in a coplin staining jar for ten minutes. 17. Rinse both sides of the slide with distilled water to remove all traces of excess gold chloride. 18. Flood the slides with oxalic acid and allow this reagent to act for ten minutes. 19. Rinse in distilled water. 20. Flood the sections with the sodium thiosulphate solution changing the solution every time it becomes turbid over a period of ten minutes. 21. Wash well in running water; then drain. 22. Dehydrate in ascending grades of alcohol, clear in xylol and mount. 142 SECTION TWO Results: Collagen is stained a reddish purple, while reticulum appears as black threads. LORRAIN - SMITH - DIETRICH STAIN For lipoids Solutions required: A. Potass, dichromate 5% aqueous. Haematoxylin (Kultschitzky) B. Haematoxylin 10% in absolute alcohol (ripened three months or longer) . . . . . . 10 ml. Acetic acid 2% aqueous . . 90 ml. C. Potass, ferricyanide .. .. 2-5 gm. Borax 2% aqueous . . . . 100 ml. Technique: Material is fixed in 10% formalin and frozen sections are em- ployed. 1. Mordant sections twenty-four to forty-eight hours in Solu- tion A at 37° C. ; then wash thoroughly in distilled water. 2. Immerse in Solution B at 37° C. for four to five hours; then wash in distilled water. 3. Differentiate overnight in Solution C. 4. Wash thoroughly in distilled water; drain and mount in Aquamount or in Farrant. Results: Lipoid substances, blue-black. LUGOL'S IODINE For the identification of glycogen in tissues Solution required: Lugol's iodine. 143 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1 . Thin slices of tissue are fixed in absolute alcohol ; then de- hydrated; cleared, and embedded in paraffin wax in the usual manner. 2. Float sections on slides with 70% alcohol and flatten by warming gently, on a warm surface (but not with a direct flame). 3. Remove excess 70% alcohol by blotting very carefully but thoroughly. 4. Treat with xylol ; then with absolute alcohol. 5. Stain in Lugol's iodine solution for ten minutes; then pour off" excess stain and carefully blot the preparation thoroughly dry. 6. Clear and differentiate with origanum oil, controlling by examination under the microscope. 7. Mount in origanum balsam. Results: Glycogen, reddish brown; tissue constituents, pale yellow. LUXOL FAST BLUE - CRESYL FAST VIOLET (Kliiver and Barrera's Stain) For the combined staining of cells and fibres in the nervous system, obviating the need for chromate treatment and haematoxylin Solutions required: A. Luxol fast blue o-i% in 95% alcohol Acetic acid 10% aqueous . . 100 ml. 0-5 ml. B. Lithium carbonate 1% aqueous . . Distilled water 5 rnl- 95 ml. C. Cresyl fast violet, CNS o-i to 0-25% aqueous Acetic acid i % aqueous 120 ml. I ml. D. Xylol Terpinol I volume 3 volumes 144 SECTION TWO Technique: Material should be fixed in io% formalin. Paraffin or frozen sections give somewhat better results than Celloidin. Affixed Celloidin give better results than loose Cel- loidin sections. (a) Frozen sections 1. Cut sections 25^11 in thickness and place them in distilled water. 2. Immerse in 70% alcohol for ten to fifteen minutes. 3. Stain from five to twenty-four, but preferably not less than sixteen hours, in the Luxol fast blue solution, in a stoppered jar in an oven at 40° C. Note: For staining four sections of the brain stem of a monkey, for example, 20 to 25 ml. of the stain should be used and then discarded. 4. Immerse in 95% alcohol and wash off the excess stain. 5. Wash in distilled water. 6. Immerse for two or three seconds, but no longer, in the lithium carbonate solution, as the first stage of differentiation. 7. Continue the differentiation in several changes of 70% alcohol until the grey and white matter can be distinguished, but taking care not to over- differentiate. 8. Wash in distilled water. 9. Immerse in the lithium carbonate solution for three to five seconds, but no longer. 10. Complete the differentiation by immersing in several changes of 70% alcohol, until the white matter is stained greenish- blue in sharp contrast with the colourless grey matter. 11. Wash thoroughly in distilled water. 12. Stain for one to two minutes in the cresyl fast violet solution, which should be warmed carefully and filtered before use. 13. Wash for two or three seconds in distilled water. 14. Differentiate in several changes of 95% alcohol until colour ceases to come away from the preparation and the alcohol is no longer tinted. 15. Clear in xylol-terpineol (Solution D). 16. Clear in xylol and mount. H5 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES (b) Paraffin sections 1. Cut sections 15 to 20jLt in thickness and fix to slides. 2. Remove paraffin wax with xylol and pass through absolute alcohol. 3. Rinse with several changes of 95% alcohol. 4. Stain with the Luxol fast blue solution for five to twenty-four hours, but preferably not less than sixteen hours, at 57° C. in an oven, taking precautions to prevent the loss of alcohol through evaporation from the staining solution. 5. Proceed exactly as at Stage 4 in the technique given above for frozen sections, except at Stage 1 1 the cresyl violet should be allowed to act for six minutes. (c) Celloidin sections (loose) 1. Cut sections 15 to 30/z in thickness and place them into 75% alcohol. 2. Stain from five to twenty-four hours, but preferably not less than sixteen, in the Luxol fast blue solution at 57° C. in an oven, taking precautions to prevent the loss of alcohol by evaporation from the staining solution. 2. Proceed exactly as at Stage 4 in the technique given for frozen sections, except at Stage 1 1 the staining time for the cresyl fast violet should be increased to three minutes. (d) Celloidin sections (affixed) 1. Cut sections 15 to 30jLt in thickness, keeping the microtome knife and tissue continually flooded with 75% alcohol. 2. Place sections on slides, which have previously been smeared with glycerine albumen. 3. If necessary flatten out the sections en the slides by rolling with a piece of glass tubing half an inch in diameter and about two inches in length, or a small glass phial will serve the purpose. 4. Drop on sufficient clove oil to cover the sections and leave the oil to act for five minutes. 5. Remove the clove oil with 95% alcohol. 6. Remove the Celloidin with absolute alcohol. 7. Wash with 95% alcohol. 146 SECTION TWO 8. Proceed exactly as at Stage 2 in the technique given for frozen sections, except at Stage 2 the staining time for cresyl fast violet should be increased to six minutes. Results: Myelinated fibres are sharply contrasted greenish-blue against the reddish-coloured Nissl cells. The technique shows the Nissl picture and differentiates between the three types of glia cells: Myelin sheaths, neurons and glia nuclei are well demonstrated. Differentiation is also obtained between the three layers of medium-sized and larger blood cells, and capillary endothelium as well as mesothelial lining of Arachnoid membrane are sharply outlined. The finer fibres of the molecular layer of the cerebral cortex can be most effectively demonstrated in paraffin sections. Bacteria and pigments in nerve cells are more clearly demonstrated with this technique than with the usual Nissl stains. References : Kliiver, Heinrich and Barrera, Elizabeth (1953), J. of Neuropath and Exp. Neurology, 12, no. 4, 400-3, " A Method for the combined staining of cells and fibres in the nervous system ". Kliiver, Heinrich and Barrera, Elizabeth (1954), X of Psychology, yj, 199-223, " On the use of Azoporphin derivatives (Phthalocyanines) in the staining of nervous tissue ". Note: In the original paper it is stated that this cell-fibre stain has been employed for peripheral nerves as well as structures of the central nervous system in amphibians, birds and mammals (rat, guinea-pig, rabbit, cat, monkey, chimpanzee, gorilla and man), and that with suitable counterstains Luxol fast blue will give ex- cellent preparations of cochlea, adrenals and numerous extraneural tissues. MacCALLUM'S STAIN For influenza bacilli and Gram-positive organisms in tissues Solutions required: A. Goodpasture* s stain: Alcohol 30% Basic fuchsin Aniline oil Phenol crystals « « ■ • • • • • « 100 ml. . 0-59 gm I ml. . I gm. H7 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES B. Picric acid saturated aqueous. C. Stirling's gentian violet. D. Gram's iodine. E. Equal volumes of xylol and aniline oil. Technique: Tissues should be fixed in Helly and embedded in paraffin wax. 1. Stain for ten to thirty seconds in Solution A; then wash in tap water. 2. Differentiate for a few seconds in formaUn till the bright red colour changes to a clear deep pink ; then wash with tap water. 3. Counterstain one to five minutes in Solution B until the sec- tion appears purplish yellow to the naked eye ; then wash with tap water. 4. Differentiate in 95% alcohol until the section appears red; then wash in tap water. 5. Stain for about five minutes in Solution C; then wash in tap water. 6. Stain for one minute in Solution D ; then, without washing, blot dry. 7. Treat in Solution E until no more colour comes out. 8. Pass through two changes of xylol; then mount. Results: Gram-positive organisms, blue. Gram-negative, red. Tissues, varying shades of red and purple. MALLORY'S STAIN - HAEMATOXYLIN For the differential staining of the pancreatic islets Solutions required: A. Distilled water . . . . . . 100 ml. Sulphuric acid cone. . . . . i ml. Potassium Permanganate . . . . i gm. 148 SECTION TWO B. Harris or Ehrlich haematoxylin. C. Lithium carbonate, saturated aqueous. D. Acid Fuchsin (Mallory) o*5% aqueous. E. Phosphomolybdic Aniline Blue - Orange G (Mallory). Technique: 1. Mammalian pancreases are fixed in Bouin and afterwards washed in 80% alcohol. 2. Dehydrate, clear, and embed in paraffin wax. 3. Cut sections 4 to 5 jit in thickness. 4. Fix sections to slides: remove wax with xylol and pass through the usual grades of alcohol down to distilled water. 5. Immerse in solution A for about one half to one minute, until the sections appear uniform reddish brown in colour. 6. Rinse in distilled water. 7. Stain with Ehrlich or Harris Haematoxylin for five to ten minutes. 8. Blue in lithium carbonate solution. 9. Wash well in tap water. 10. Wash with distilled water. 11. Stain in Mallory's Acid Fuchsin, controlling under micro- scope until the A cells are red, and the B cells are pink. If overstained, wash out with distilled Vv^ater until the above effects are obtained. 12. Stain in solution E for twenty minutes to twelve hours according to the condition of the pancreas and degree of dif- ferentiation in the first stain. Results: Nuclei are stained dark violet; nucleoli, red. Cytoplasm in A cells contains red granules. Cytoplasm of B cells contains blue granules. Cytoplasm in acinar cells varies from red to pale violet with deep violet ergastoplasm. Canahcular cells, blue-grey. Connective tissue blue. Erythrocytes red. Mucus, azure blue. Reference: Isaac, J. P. and Aron, C. (1952), Bull. Mies. Appl. ser. 2, 2, 99-102. 149 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES MALLORY STAIN - HAEMATOXYLIN For differential staining of acidophils, basophils and chromophobes in mouse pituitary Solutiotis required: A. Zenker - Formal : Zenker's Fluid . . . . . . 95 ml. Formaldehyde . . . . . . 5 ml. B. Formic acid 10% aqueous. C. Iodine 0-5% in 70% alcohol. D. Sodium thiosulphate 075% in 10% alcohol. E. Harris haematoxylin. F. Lithium carbonate satd. aqueous. G. Acid fuchsin 0-5% aqueous. H. Phosphomolybdic acid 1% aqueous. I. Mallory's Aniline blue-orange G. J. Carbol xylol. Technique: 1. Pituitary gland together with bone to which it is attached is fixed in solution A for 4-8 hours, or overnight in a refrigerator. 2. Wash in running tap water for 8 to 10 hours. 3. Decalcify by immersing in 10% formic acid for 24 hours. 4. Wash in running tap water for 2 to 4 hours. 5. Dehydrate, clear and embed in paraffin wax 56 to 58° C. in the usual way. 6. Cut sections about 4)Lt in thickness and mount on slides with glycerine albumen. 7. Dry the slides thoroughly in an oven at 56° C. for one to two hours, or overnight at room temperature. 8. Remove wax with xylol; then wash with two changes of absolute alcohol. 150 SECTION TWO 9. Wash with 90% followed by 70% alcohol. 10. Immerse for half to two minutes in solution C (Iodine). 1 1 . Wash well with water. 12. Immerse in solution D (thiosulphate) for half to two minutes or until the natural colour of the sections is restored. 13. Wash well with tap water. 14. Rinse in distilled water. 15. Stain in Harris haematoxylin for 2-3 minutes. 16. Rinse in tap water. 17. Blue in the lithium carbonate solution for i minute. 18. Rinse well in distilled water. 19. Stain in the acid fuchsin solution for 1-2 minutes. 20. Rinse quickly in tap water. 21. Immerse in the phosphomolybdic acid solution for 2-5 minutes. 22. Without washing, pass the slides directly into Mallory's Aniline blue - orange G and leave therein for 1-2 hours. 23. Differentiate with 95% alcohol, controlling by examination under the microscope until the blue granules are intense, but the fuchsinophil granules are clearly visible. 24. Rinse with absolute alcohol. 25. Rinse with carbol xylol. 26. Immerse in two changes of xylol for 10 minutes in each. 27. Mount in Clearmount of D.P.X. Results: The nuclei of all cells are stained dark blue black. Granules of basophils, blue. Acidophil granules, brilliant red. Non-granular cytoplasm of chromophobes, light grey. Erythrocytes, orange to red. Bone, intense blue. Reference: Gude, William D. (1953), Stain Tech., 28, no. 3, 161-2. M 151 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES MALLORY STAIN - HAEMATOXYLIN (Ehrlich) For Negri bodies in sections of brain Solutions required: A. Ehrlich haematoxyhn. B. Orange G. . . . . . . . . 0-5 gm. Phosphotungstic acid, saturated aqueous . . . . . . . . 100 ml. C. Acid fuchsin . . . . , . 0-5 gm. Phosphotungstic acid . . . . 0-5 gm. Acetic acid 1% . . . . . . 100 ml. D. Phosphotungstic acid . . . . 2 gm. Phosphomolybdic acid . . . . 2 gm. Picric acid, saturated, aqueous . . 70 ml. Absolute alcohol . . . . • • 30 ml. E. Aniline blue, aqueous . . . . i gm. Distilled water . . . . . . 98 ml. Glacial acetic acid . . . . . . 2 ml. Technique: 1 . Fix paraffin sections to slides ; dewax and take down to dis- tilled water in the usual way. 2. Stain in Ehrlich haematoxyhn for five minutes. 3. Blue in tap water, or in lithium carbonate solution, for two minutes. 4. Rinse in distilled water. 5. Stain for one minute in the orange G solution. 6. Wash in tap water until only the erythrocytes are stained yellow. 7. Rinse in distilled water. 8. Stain for ten minutes in the acid fuchsin solution. 9. Rinse in distilled water. 10. Differentiate for five minutes in solution D. 1 1 . Rinse in distilled water. 152 SECTION TWO 12. Rinse in i% acetic acid. 13. Dehydrate, clear and mount. Results: Negri bodies, purplish red with blue granulations. Cytoplasm of neurones, bluish. Nucleoli, dark purple. Erythrocytes, yellow. Reference: Zlotnik, I. (1953), Nature, 172, no. 4386, 962-3. MALLORY HEIDENHAIN STAIN (Jane E. Cason's modification) A rapid one-step method for connective tissue Solutions required: Phosphotungstic acid crystals A. R. i gm. Orange G. . . . . . . 2 gm. Aniline blue, water soluble . . i gm. Acid fuchsin . . . . • • 3 gni. Distilled water . . . . . . 200 ml. Technique: I . Fix pieces of tissue in Zenker-formol for preference, although Bouin's fluid, formalin and alcohol have been used with success. a. Embed in paraffin wax and cut sections 6// in thickness. 3. Fix sections to slides and remove wax with xylol. 4. Pass through descending grades of alcohol and if Zenker- formol has been used as the fixative, treat with iodine and sodium thiosulphate as usual to remove mercurial precipitate. 5. Take down to tap water. 6. Immerse for five minutes in the staining solution. 7. Wash in running tap water for three to five minutes. 8. Dehydrate rapidly through the usual graded alcohols. - 9. Clear in xylol and mount. 153 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Results: Appear to be the same as those listed by Mallory (1938), i.e. collageneous fibrils, intense blue. Ground-substance of cartilage and bone, mucus, amyloid, and certain other hyaline substances are stained in varying shades of blue. Nuclei, fibroglia, myoglia and neuroglia fibrils, nucleoli, axis cylinders and fibrin are stained red. Erythrocytes and myelin, yellow. Elastic fibrils are stained pale pink or yellow. Abstract: Cason, Jane E., Stain Technology (1950), 25, No. 4, 225-6. MALLORY'S PHOSPHOTUNGSTIC ACID HAEMATOXYLIN A general stain for vertebrate tissues Solutions required: Haematoxylin 10% in absolute alcohol (ripened for three months or longer) . . . . i ml. Phosphotungstic acid . . . . 2 gm. Distilled water . . . . . . 100 ml. Note: If ripened haematoxylin solution is not available, the following artificially ripened stain should be used: Haematoxylin (dry) o-i gm., phosphotungstic acid 2 gm., distilled water 100 ml., potassium perman- ganate 1% aqueous 1-77 ml. Technique: 1 . Fix in Zenker. Embed in paraffin wax. 2. Bring sections down to distilled water. 3. Treat with iodine to remove mercuric percipitate. 4. Remove iodine with 0-5% aqueous sodium hyposulphite. 5. Wash thoroughly in running water. 6. Immerse for five to ten minutes in 0-25% potass, perman- ganate ; then wash in tap water. 7. Immerse for ten to twenty minutes in 5% oxalic acid; then wash thoroughly with tap water. 154 SECTION TWO 8. Stain twelve to twenty-four hours in haematoxylin solution, prepared as above. 9. Wash in tap water; dehydrate with 95% and absolute alcohol. 10. Clear in xylol and mount. Results: Nuclei, centrioles, achromatic spindles, fibroglia, myoglia, neuroglia fibrils, fibrin, contractile elements of striated muscle, blue. Collagen, reticulum, ground substances of cartilage and bone, yellowish to brownish red. Coarse elastic fibrils, faint purple. MARSHALL RED - VICTORIA GREEN A general stain, particularly suitable for class work Solutions required: A. Marshall red, saturated aqueous. B. Victoria green saturated in 70% alcohol. Technique: 1. Fix sections to slides; dewax and take down to distilled water in the usual manner. 2. Stain in the Marshall red solution for twenty minutes. 3. Rinse in distilled water. 4. Stain in the Victoria green solution for half an hour, 5. Rinse in 70% alcohol followed by 90% alcohol. 6. Dehydrate with absolute alcohol. 7. Clear in xylol and mount. Results: Myofibrils, sage green. Nuclei, bright carmine. The results vary somewhat, but the muscle fibres always appear greenish to greenish -grey, while the nuclei are red. White matter of spinal cord, yellowish-green. Cartilage, pink. Retina stands out well as the rods and cones appear bright bluish-green. Erythrocytes unstained. Reference: Cannon, H. G. (1941), J. Roy. Micro. Soc.y series III, 61, parts 3 and 4. MEDICAL AND BIOLOGICAL STAINING TECHNIQUES MASSON'S TRICHROME STAIN For connective tissues Solutions required: A. Iron alum 5% aqueous. B. Regaud's haematoxylin solution. C. Picric acid, saturated in 95% alcohol . . . . . . . . 20 ml. Alcohol 95% . . . . . . 10 ml. D. Ponceau fuchsin. E. Phosphomolybdic acid 1% aqueous. F. Aniline Blue 5% in 2% acetic acid. Technique: 1. Fix pieces of tissue in Bouin's fluid for three days or in Regaud's fluid for one day. 2. Wash in running water; dehydrate; clear and embed in parafiin wax as usual. 3. Sections 5/1 in thickness are fixed to slides; de-waxed and passed through descending grades of alcohol down to distilled water in the usual manner. 4. Mordant in Solution A for five minutes at 45° C. to 50° C. 5. Wash well in distilled water. 6. Stain for five minutes in Regaud's haematoxylin at 45° C. to 50° C. 7. Rinse in distilled water. 8. DiflFerentiate in picric alcohol (Solution C above) controlling by examination under the microscope, while the preparation is still wet. 9. Wash in running tap water for a minute or so. 10. Stain for five minutes in the Ponceau fuchsin solution. 11. Rinse in distilled water. 12. Diff"erentiate in the phosphomolybdic acid solution for five minutes. 156 SECTION TWO 13. Add 0-5 ml. of the acetic aniline blue (Solution F above) to the phosphomolybdic acid on the slide and mix by rocking the slide gently. Allow this mixture to act for five minutes. 14. Pour off excess liquid and rinse in distilled water. 15. Immerse in phosphomolybdic acid solution again, for five minutes. 16. Transfer to i % acetic acid and leave therein for five minutes. 17. Wash in distilled water. 18. Dehydrate in 95% alcohol, followed by absolute alcohol; clear in xylol ; mount. Results: Collagen, deep blue. Neuroglia fibrils, red. Nuclei, black. Argentaffin granules, black or red. METHYL BLUE - EOSIN (Mann) For demonstrating the various types of cells in the anterior lobe of the pituitary and for the study of the relationship and development of the blood vessels Solution required: Methyl Blue-Eosin (Mann's stain). Technique: 1 . Paraffin sections of tissues which have been fixed in a fluid con- taining mercuric chloride are mounted on slides and treated by the standard technique for the removal of mercuric precipitate (see page 28). 2. Bring the sections down to distilled water; then stain for a quarter of an hour to two hours (the longer time is required if it is desired to demonstrate anterior lobe of pituitary). 3. Wash and differentiate in tap water. 4. Dehydrate rapidly with two changes of absolute alcohol. 5. Clear in xylol ; mount in xylol balsam and examine under the low power, as the stain is too diffuse for critical high power work. 157 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Results: Nuclei are stained blue; karyosomes, dark blue; plasmosomes, red; basophil cytoplasm, blue; oxyphil cytoplasm and oxyphil granules, red. METHYL GREEN For amyloid Solution required: Methyl Green i% aqueous. Technique: 1. Fix tissues in absolute alcohol or in io% formalin and cut frozen sections. 2. Immerse in the methyl green solution for five to ten hours. 3. Wash well in distilled water. 4. Mount in neutral glycerine, or in aquamount. Results: Amyloid is stained reddish violet, while other tissue elements are green. METHYL GREEN - PYRONm (Pappenheim-Unna) For plasma cells Solution required: Methyl Green - Pyronin (Pappenheim-Unna). Technique: 1. Paraffin sections of absolute alcohol-fixed tissues are mounted on slides and brought down to distilled water by the standard method. 2. Stain in Methyl Green - Pyronin (Pappenheim-Unna) from five to thirty minutes. 3. Rinse in distilled water; drain and blot carefully. 4. Dehydrate rapidly in two changes of absolute alcohol. 5. Clear in xylol; and mount. 158 SECTION TWO Results: Plasmosomes and oxyphil constituents of cytoplasm are stained red; chromatin reticulum and karyosomes, bluish green; cyto- plasm of plasma cells, brilliant red ; ground-substance of hyaline cartilage, yellowish red; bacteria, vivid red. METHYL VIOLET 6B For amyloid Solution required: Methyl Violet 6B aqueous i%. Technique: 1. Material should be fixed in alcohol and embedded in paraffin wax (although frozen sections may be employed). 2. Paraffin sections are fixed to slides and brought down to dis- tilled water. 3. Stain for two to five minutes in the methyl violet 6B solution. 4. Rinse quickly in distilled water. 5. Differentiate with 1% acetic acid, controlling by examination under the microscope, until the amyloid appears reddish in colour. 6. Wash in distilled water. 7. Mount in glycerine jelly. Results: Amyloid substance is stained red, while cells and nuclei are in varying shades of blue. Notes: (a) The technique is not suitable for material which has under- gone prolonged fixation in formalin. {h) The stain is extracted from the amyloid by alcohol and for this reason glycerine jelly is used as the mountant, thereby obvi- ating the use of alcohol for dehydration. 159 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES METHYL VIOLET - METANIL YELLOW For typhus fever rickettsiae in lungs of mice Solutions required: A. Methyl violet loB . . . . . . o-oi% aqueous. B. Distilled water . . . . . . 90 ml. Acetic acid I % .. .. .. 10 ml. C. Metanil yellow . . . . . . o-oi% aqueous. Technique : 1. Fix pieces of lung in 10% neutral formalin. 2. Dehydrate, clear and embed in paraffin wax in the usual manner. 3. Fix sections to slides and take down to distilled water as usual. 4. Stain in the methyl violet solution for half to one hour. 5. Differentiate in solution B, controlling by examination under the microscope until the cytoplasm is decolorized. 6. Counterstain in the metanil yellow solution for a few seconds. 7. Dehydrate in acetone. 8. Mount in D.P.X. or Clearmount. Results: Rickettsiae are stained violet. Reference: Nyka, W. J. (1945), J^. Path, and Bad., 52, 317-24. METHYL VIOLET - PYRONIN - ORANGE G (Bonney's Triple Stain) For chromatin, connective tissue, keratin Solutions required: A. Methyl violet 6B (Jensen) 1% aqueous . . . . . . 25 ml. Pyronin 10% aqueous . . . . 10 ml. Distilled water . . . . . • 65 ml. 160 SECTION TWO B. Acetone . . . . . . . . loo ml. Orange G aqueous 2% . . . . About lo ml. Add orange G solution drop by drop to the ace- tone, with shaking, until the flocculent precipitate formed just redissolves with further addition of orange G solution. Technique: 1. Small pieces of tissue are fixed in acetic-alcohol or in mer- . curie chloride. 2. Wash; dehydrate; clear; embed in paraffin wax. 3. If mercuric chloride has been used for fixation treat sections for the removal of mercuric precipitate by the standard method. 4. Take sections down to distilled water. 5. Immerse for two minutes in the methyl violet pyronin (Solution A above). 6. Pour off excess stain and carefully wipe the slide dry. 7. Flood the preparation with acetone-orange G solution. 8. Pour off after a few seconds. 9. Flood the preparation with a fresh lot of acetone-orange G solution and pour off after a few seconds. 10. Wash quickly in pure acetone. 1 1 . Rinse with two lots of xylol. 12. Mount in balsam. Results: Cytoplasm, red; chromatin, violet; keratin, violet; connective tissue, yellow. METHYLENE BLUE - BASIC FUCHSIN Rapid method of demonstrating. Negri bodies in sections Solutions re 'quired: A. Methylene blue . . Basic fuchsin Methyl alcohol, pure Glycerine, pure . . m • • • • • • • . . I gm. .. 1-75 gm 100 ml. 100 ml. 161 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES B. Potass, hydroxide N.20 . . . . 6-25 ml. Distilled water . . . . . . 93-75 ml. C. Solution A . . . . . . 10 ml. Solution B . . . . . . 0-25 ml. Note: Solution C should be freshly prepared as required. Technique: 1. Blocks not exceeding 3 mm. in thickness of fresh tissue from the hippocampus major and cerebellum should be fixed in Zenker for twelve to twenty-four hours. 2. Remove mercurial precipitate with iodine in alcohol by the usual method (see page 28) ; then wash in running water for three to six hours. 3. Dehydrate in dioxane (see page 36) and embed in paraffin wax. 4. Sections not more than 4/x thick are mounted on slides and brought down to distilled water. 5. Flood slides with Solution C and steam gently for five min- utes ; then cool and wash rapidly in tap water. 6. Decolorize and differentiate each section separately by waving the slide gently in a jar of 90% alcohol until the section assumes a faint violet colour. 7. Dehydrate rapidly in 95% and absolute alcohol; clear in xylol and mount. Results: Negri bodies, deep red ; granular inclusions, dark blue ; nucle- oli, bluish black; cytoplasm, bluish violet; erythrocytes, dull reddish brown. METHYLENE BLUE - BASIC FUCHSIN For rickettsia in sections Solutions required: A. Methylene blue 1% aqueous. B. Basic fuchsin 0-5% aqueous. C. Citric acid 0*5% aqueous. 162 SECTION TWO Technique: 1. Tissues are fixed in Regaud's fluid, washed in running water, dehydrated, cleared and embedded in paraffin wax as usual. 2. Fix sections to slides; de-wax; pass through the alcohol down to distilled water in the usual manner. 3. Stain in the methylene blue solution for twelve to sixteen hours. 4. Decolorize with 95% alcohol. 5. Counterstain with the basic fuchsin solution for fifteen min- utes. 6. Decolorize for one to three seconds in the citric acid solu- tion. 7. Pour off excess citric acid ; rinse in distilled water; drain and blot carefully. 8. Decolorize and dehydrate rapidly in absolute alcohol. 9. Clear in xylol and mount in dammar xylol. Results: Rickettsia are stained a deep red; surrounding tissue, pink; background, light blue. METHYLENE BLUE POLYCHROME (Unna) For mast cells in sections Solutions required: Methylene blue, polychrome (Unna) . . . . . . 100 ml. Potash alum . . . . • • 5 gm. Technique: 1. Material is fixed in absolute alcohol and embedded in Celloidin. 2. Stain sections in a watch glass from three to sixteen hours in the methylene blue solution. 3. Rinse well in distilled water. 163 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 4. Dehydrate with 95% alcohol. 5. Clear in origanum oil. 6. Mount in Canada balsm or in Cristalite. Results: Mast cell granules are stained red ; nuclei, blue. METHYLENE BLUE POLYCHROME - GLYCERINE ETHER (Unna) For differentiating mast cells and plasma cells Solutions required: A. Formalin (40% formaldehyde) . . 50 ml. Absolute alcohol . . . . 100 ml. B. Polychrome methylene blue (Unna) C. Glycerine ether (Unna) . . • • 5 nil- Distilled water . . . . . . 35 ml. Technique: 1. Fix tissues in Solution A; dehydrate; clear; embed in paraffin wax. 2. Immerse sections in the methylene blue solution for ten minutes. 3. Rinse in distilled water. 4. Differentiate in the glycerine ether mixture from one half to one minute until the sections appear to be deep sky blue to the naked eye (care should be taken that the sections are not over- differentiated). 5. Wash thoroughly in distilled water for a few minutes. 6. Fix sections to slides and carefully blot with filter paper. 7. Dehydrate rapidly with absolute alcohol. 8. Clear in xylol and mount. Results: Nuclei are stained blue, while mast cells are red and plasma are blue. 164 SECTION TWO MUCICARMINE - METANIL YELLOW - HAEMATOXYLIN For mucin and connective tissue Solutions required: A. Haematoxylin (Weigert) A. B. Haematoxylin (Weigert) B. C. Metanil yellow 0-25% aqueous. D. Mucicarmine (Southgate). Technique: 1. Fix material in 10% formalin. 2. Dehydrate, clear; embed in paraffin wax. 3. Fix sections to slides; de-wax with xylol and pass through the usual descending grades of alcohol. 4. Rinse with distilled water. 5. Stain sections for one minute in a freshly prepared mixture consisting of equal volumes of Solution A and B. 6. Wash in distilled water. 7. Immerse in Solution C for about two minutes. 8. Rinse quickly with distilled water. 9. Immerse in the mucicarmine solution for forty-five minutes. 10. Rinse quickly with distilled water. 11. Rinse quickly with 95% alcohol. 12. Dehydrate rapidly but thoroughly with absolute alcohol. 13. Clear in xylol and mount. Results: Mucin is stained red, while connective tissue is yellow, and nuclei are black. 1 6s MEDICAL AND BIOLOGICAL STAINING TECHNIQUES MUCICARMINE (Mayer) Solutions required: Mucicarmine, stock solution . . i volume Alcohol 70% . . . . . . 10 volumes Note: This diluted solution should be freshly prepared. Technique: Tissues should be fixed in absolute alcohol for five to eight hours and embedded in paraffin wax, Celloidin or L.V.N. 1. Bring paraffin sections down to distilled water; then stain for ten to twenty-five minutes in the above solution. 2. Wash rapidly with distilled water. 3. Dehydrate with 70%, 95% and absolute alcohol. 4. Paraffin sections are cleared in xylol; Celloidin or L.V.N. sections are cleared in terpineol or origanum oil. Results: Mucin is stained red. MUCICARMINE (Southgate) This is used in accordance with the Mayer technique, but i vol- ume of the stock solution is diluted with 9 volumes of distilled water instead of 10 volumes 70% alcohol. Southgate's modification gives more uniform results than Muci- carmine prepared by Mayer's original formula. MUCIHAEMATEIN I For mucus Solution required: Haematein Aluminium chloride Glycerine, pure . . Distilled water . . 0-2 gm . . o-i gm . . 40 ml. . . 60 ml. 166 SECTION TWO Technique: 1. Material is fixed in absolute alcohol; cleared; and embedded in paraffin wax. 2. Sections are stained for ten minutes in the above solution. 3. Wash with distilled water. 4. Dehydrate by plunging the slide into two or three changes of 95% alcohol. 5. Pass through absolute alcohol ; then clear in xylol ; mount. Results: Mucus is stained blue, while the remainder is colourless. NADI REACTION For oxidase granules Solutions required: A. a-Naphthol . . . . . . i gm. Distilled water . . . . . . 100 ml. Place in a 250-ml. flask and boil until the a-Naph- thol begins to melt; then add 40% potassium hy- droxide aqueous solution drop by drop until the solution becomes yellowish-blue in colour and the a-Naphthol is still not completely dissolved. B. Cold tap water . . . . . . 100 ml. *D imethy 1 -p -pheny lenediamine base . . . . . . . . 0-5 gm. Place the water in a clean amber bottle, open the ampoule by filing a groove at one end then breaking in the usual manner. Tip the contents of the am- poule into the bottle; then replace the stopper and allow the bottle to stand, with occasional shaking, for twenty-four hours. Care should be taken that the Dimethyl-p-phenylenediamine base does not come into contact with the body. This solution deteriorates after three or four weeks. *Note: This should be purchased in a sealed ampoule. N 167 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES C. Gram's iodine solution. D. Carmalum. Technique: 1. Thin pieces of fresh tissue (not more than 3 mm. thick) are fixed for three to five hours in formalin-saUne solution. 2. Make frozen sections and collect them in distilled water. 3. Immerse in a mixture consisting of equal volumes of Solu- tions A and B (Note: This mixture must be made and filtered immediately before use) for about five minutes until the sections turn blue. 4. Rinse sections rapidly in distilled water. 5. Immerse in Gram's iodine solution until the sections turn brown. 6. Transfer to distilled water to which two drops of lithium car- bonate 0*5% aqueous have been added for each 100 ml. of distilled water, for a quarter to twenty-four hours until the sections have regained their blue colour. 7. Wash in distilled water then counterstain in carmalum for two to five minutes. 8. Mount in glycerine jelly or Apathy's medium or in Aqua- mount. Results: Oxidase granules are stained blue while nuclei are pink. Some- times fat is stained also. NAPHTHOL BLUE BLACK - HAEMATOXYLIN - BRILLIANT PURPURIN AZOFUCHSIN For collagen, reticulum, smooth muscle, etc. Solutions required: A. Weigert's Haematoxylin A. B. Weigert's Haematoxylin B. C. Brilliant purpurin R. (C.I. No. 454) in 1% acetic acid . . . . . . 30 ml. Azofuchsin 1% in 1% acetic acid. . 20 ml. 168 SECTION TWO D. Naphthol blue black (C.I. No. 246) i gm. Picric acid, satd., aqueous . . 100 ml. Technique: 1. Paraffin sections are fixed to slides and taken down to 70% alcohol in the usual manner. 2. Stain for six minutes in a freshly prepared mixture consisting of equal parts of solutions of A and B. 3. Wash in tap water. 4. Counterstain for five minutes in solution C. 5. Wash in 1% acetic acid. 6. Stain in solution D for five minutes. 7. Rinse in 1% acetic acid for two minutes. 8. Pass through the usual ascending grades of alcohol dehydrate in absolute. 9. Clear in Xylol. 10. Mount in D.P.X. or Clearmount. Results: Collagen, reticulum and basement membranes, dark green. Smooth muscle, brown. Nuclei, brownish black. Reference: Lillie, R. D. (1945), J. Tech. Meth., 25, 47. NAPHTHOL GREEN B - HAEMATOXYLE^ (Weigert) For connective tissue Solutions required: A. Weigert's haematoxylin, A. B. Weigert's haematoxylin, B. C. Eosin, yellowish, 1% in tap water. D. Ferric chloride, hydrated 10%. E. Naphthol Green B, 1% aqueous. F. Equal volumes of acetone and xylol. 169 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Paraffin sections are mounted on the slide and brought down to distilled water in the usual manner. 2. Stain for six minutes in a freshly prepared mixture consisting of equal volumes of Weigert's haematoxylin A and B. 3. Wash thoroughly in tap water; then stain for three minutes in the eosin solution. 4. Wash in tap water ; then immerse in the ferric chloride solu- tion for five minutes. 5. Rinse well in distilled water; then stain for five minutes in the naphthol green solution. 6. Differentiate for two or three minutes in 1% acetic acid. 7. Drain well; then dehydrate with acetone, afterwards clear- ing in acetone-xylol (as above); then mount. Results: Connective tissue, green; muscle and cytoplasm, pink. Reference: Lillie, R. D. (1945), J^. Tech. Meth., 25, 32. NEUTRAL RED - FAST GREEN For staining both Gram-positive and Gram-negative bacteria in sections Solutions required: A. Aniline crystal violet. B. Gram's iodine. C. Absolute alcohol . . . . 98 ml. Glacial acetic acid . . . . 2 ml. D. Twort's stain, modified (neutral red-fast green) Technique: 1. Fix tissue in 5% formal-saline, dehydrate, clear; embed in paraffin wax. Cut sections 3^^ in thickness. 2. Stain in aniline crystal violet for three to five minutes. 170 SECTION TWO 3. Pour off excess and blot, without washing. 4. Flood with Gram's iodine and allow the stain to act for three minutes. 5. Destain with the acetic acid alcohol (Solution C above) until no more colour comes away, and the sections assume a dirty straw colour. 6. Rinse quickly in distilled water. 7. Stain in neutral red-fast green diluted one part with three parts of distilled water, for five minutes. 8. Wash quickly with distilled water. 9. Decolorize with the acetic alcohol solution until no more red stain comes out. 10. Rinse quickly in absolute alcohol. 1 1 . Clear in xylol and mount. Results: Nuclei are stained red, while cytoplasm is light green. Gram- positive bacteria, dark blue. Gram-negative bacteria, pink. Erythrocytes, green. J. Path. & Bad., 59, 357-8, Ollett, W. S. (1947). NILE BLUE SULPHATE For demonstrating fatty acids and neutral fats Solution required: To 100 ml. saturated aqueous Nile Blue sulphate add 0'5 ml. cone, sulphuric acid; then boil under a reflux condenser for two hours ; allow to cool ; then use as follows : Technique: 1. Fix small pieces of tissue in 10% formalin. 2. Frozen sections are stained for about ten minutes to half an hour at 37° C. ; or overnight at room temperature. 171 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 3. Differentiate in 2% acetic acid, 4. Rinse in distilled water; mount in Farrant. Results: Free fatty acids, blue. Neutral fats, red. NILE BLUE - PICRO FUCHSIN (Murray-Drew) For bacteria and actinomyces in pathological tissues Solutions required: A. Picro fuchsin (Van Gieson). B. Nile Blue sulphate aqueous 1%. Technique: 1. Formalin-fixed material is embedded in paraffin wax, or frozen sections may be employed. 2. Take sections down to distilled water in the usual manner then stain in picro fuchsin for two or three minutes. 3. Wash with distilled water. 4. Stain in Nile Blue sulphate for four to twenty-four hours. 5. Rinse in distilled water until the washings are tinted pale blue. 6. Drain off excess water; then blot the preparation carefully but thoroughly. 7. Dehydrate rapidly in absolute alcohol; then clear in xylol. 8. Differentiate in clove oil for five to fifteen minutes (for para- ffin sections) or for several hours in the case of thick frozen sections. 9. Rinse in two or three changes of xylol to remove clove oil; then mount. Results: Bacteria and chromatin are stained blue; collagen fibres, red; mast cell granules, blue-black; fibrin, blue or reddish orange; erythrocytes and keratin, yellow. 172 SECTION TWO I ORANGE G - CRYSTAL VIOLET (Bensley) For secretion antecedents of serous or zymogenic cells. This stain is particularly suitable for sections of the stomach or pancreas Solutions required: A. Osmic acid 2% . . . . • • 4 ml. Potassium dichromate 5% . . 4 ml. Glacial acetic acid . . . . i drop Distilled water . . . . 2 ml. B. Neutral gentian Orange G 8% aqueous . . • . 25 ml. Crystal violet 4% aqueous . . 25 ml. Mix thoroughly by shaking until almost complete precipitation takes place ; then allow the preparation to stand for an hour, afterwards collecting the preci- pitate on a filter. Wash the precipitate on the filter with about 250 ml. distilled water; then dry and dis- solve it in 25 ml. absolute alcohol. C. Solution B (as above) . . . . 10 ml. Alcohol 20% . . . . Sufficient to impart a rich port-wine colour. Allow to stand for twenty-four hours; then filter. D. Absolute alcohol . . . . i volume Clove oil . . . . . . • • 3 volumes Technique : 1. Tissues are fixed in Solution A (as above) for twenty-four hours, and afterwards embedded in paraffin wax in the usual man- ner. 2. Stain sections for twenty-four hours in Solution C (as above) ; then pour off excess stain, and blot the preparation carefully. 3. Dehydrate by immersing in two or three changes of acetone. 4. Clear in xylol; then differentiate in Solution D (as above), controlling under the microscope. 5. Rinse in two changes of xylol; then mount in balsam. 173 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Results: Zymogen granules, violet; granules of acidophil cells are stained orange-red, while those of basophil cells are violet ; back- ground, brown. ORCEIN For elastic fibres and connective tissue Solutions required: A. Orcein . . . . . . . . i gm. 70% alcohol . . . . . . 100 ml. Heat on a water bath to dissolve; cool; filter; then add : Hydrochloric acid, cone. . . i ml. Shake well. B. Unna's polychrome methylene blue. This should be diluted i : 10 or i : 15 for use. 1. Fix tissues (any fixative may be employed); dehydrate; clear; and embed. 2. Paraffin sections are brought down to 70% alcohol in the usual manner; flood with freshly filtered orcein solution, pre- pared as above, and warm gently for ten to fifteen minutes, until the solution thickens. Alternatively, the sections may be stained overnight at room temperature. 3. Wash thoroughly with 70% alcohol. 4. Wash thoroughly with distilled water. 5. Stain in diluted polychrome methylene blue until the nuclei are bright blue to blue-black; the time necessary may be ascer- tained by examining under the microscope at intervals. 6. Differentiate in 95% alcohol, followed by absolute. 7. Clear in xylol and mount. Results: Elastic fibres, deep brown. Nuclei, bright blue or blue to black. Connective tissue, pale brown. 174 I SECTION TWO Note: If it is desired only to stain the elastic fibres, omit No. 5, and treat sections with acid alcohol for a few seconds before washing with distilled water. ORCEIN - ANILINE BLUE - ORANGE G A differential stain for elastic fibres, collagen, keratin, etc. I gm. 100 ml. 0-6 ml. 49 ml. 0*5 ml. 0-5 gm. 2gm. 100 ml. Solutions required: A. Orcein Alcohol 70% Hydrochloric acid, cone. . . B. Alcohol 50% Hydrochloric acid, cone. . . C. Mallory's Aniline Blue - Orange G. Aniline blue, aqueous Orange G Phosphomolybdic acid 1%.. D. Orange G 1% in absolute alcohol. Technique: 1 . Fix material in Bouin and embed in paraffin wax. 2. Sections, about 8/x in thickness, are fixed to slides, dewaxed with xylol and passed through the usual descending grades of alcohol to distilled water. 3. Stain for one and a half hours in the orcein solution in a closed staining jar. 4. Differentiate with solution B, controlling under the micro- scope, until most of the pink is extracted from the sections. Note : The duration of the differentiation will vary according to the nature of the material and to the thickness of the sections. 5. Wash thoroughly with running tap water. 6. Wash with distilled water. 7. Immerse in solution C diluted with an equal volume of distilled water, for one to two minutes. 8. Rinse with 95% alcohol. 9. Rinse with two lots of solution D. 175 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 10. Rinse quickly with absolute alcohol. 1 1 . Clear in xylol and mount. Results: Elastic fibres, red. Collagen, blue. Muscle fibres, pale orange to dirty yellow. Cytoplasm, varying shades of yellow. Erythro- cytes, golden yellow. Kerantinized material, bright yellow. 1 Reference: Margolena, L. A. and Dolnick, E. H., Stain Tech., (195 1), 26, 1 19-21. ORCEIN - ANILINE SAFRANIN For elastic and connective tissue fibres Solutions required: A. Orcein (Unna) | to 1% in 80% alcohol . . . . . . . . 100 ml. Hydrochloric acid, cone. . . i ml. B. Safranin O, water soluble . . i gm. Aniline water . . . . . . 48 ml. Absolute alcohol . . . . 52 ml. Technique: 1. Sections are mounted on slides and brought down to 90% alcohol in the usual manner. (If the tissues have been fixed in a fluid containing mercury, the mercurial deposit is removed by the standard technique ; the sections are then taken from 70% to 90% alcohol, then direct into the orcein stain.) 2. Stain from twenty to sixty minutes with the orcein solution in a stoppered staining jar. 3. Rinse in acid alcohol until stain ceases to come out of the preparation. 4. Rinse in 70% alcohol; then in distilled water. 5. Stain in aniline safranin (Solution B) for five minutes; then rinse in water. 176 SECTION TWO 6. Differentiate by dipping into 90% alcohol then examining rapidly under the microscope. Repeat this process until the cell nuclei are well brought out, stained clear bright red. 7. Dehydrate by rinsing quickly in absolute alcohol ; then clear in xylol and mount in balsam. Results: Elastic fibres are stained dark to reddish brown; cell nuclei, bright red; ground substance of hyaline cartilage, yellow. ORCEIN - GIEMSA STAIN For sjrphilitic tissue, particularly dermatological specimens Solutions required: A. Orcein (Unna-Tanzer) : Orcein Alcohol 70% HCl, concentrated B. Absolute alcohol HCl, concentrated 0-5 gm. 100 ml. 0*6 ml. 100 ml. 0'5 ml. C. Giemsa stain . . . . . . 0-25 ml. Distilled water . . . . . . 100 ml. Note: Solution C should be prepared freshly, as required, from Giemsa stain. D. Absolute alcohol . . . . 100 ml. Eosin 1% alcoholic . . . . i ml. Technique: 1. Paraffin sections are brought down to 70% alcohol. 2. Stain for one half to one hour in Solution A ; then rinse for two to five minutes in distilled water. 3. Wipe off excess water; dip into 95% alcohol for a few sec- onds; then decolorize with absolute alcohol for five to twenty- 177 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES five minutes, or until the sections assume a pale brown colour and the elastic fibres stand out, deep purple to black, under the low- power objective. 4. Decolorize in Solution B until the background is almost colourless. This usually takes two to seven minutes. Note: Decolorization must not be extended more than ten minutes, as otherwise the thin elastic fibres will become destained. 5. Immerse in tap water for five to ten minutes. 6. Stain for two to twelve hours with Solution C until the epithelial and other cells are deep blue ; connective tissue, greyish pink or greyish blue or blue. 7. Wipe off excess stain and dehydrate and decolorize in Solution D, controlling under the microscope. Note: Decolorization must be stopped when the connective tissue has lost all trace of blue and has assumed a rose tint. The blue tinge is removed fairly rapidly in Solution D. The epidermis should remain bright blue. 8. Immerse in two changes of absolute alcohol for two minutes in each. Clear in xylol, and mount. Results: Nuclei, deep blue. Cytoplasm of the epidermis, muscle cells and connective tissue cells, light blue. Plasma cells, dark greyish blue. Eosinophilic granules, bright red. Mast cell granules, meta- chromatic (varying shades of) purple. Neutrophilic granules, only faintly stained. Erythrocytes, reddish brown. Collagenous fibres, pale rose to brownish pink. Elastic fibres, dark brown to black. Senile degenerated connective tissue (collacin, elacin and collastin), various shades of dark grey and blue. Cartilage, metachromatic (varying shades of) purple. Decalcified bone, light brown. Keratin, blue (poorly stained). Stratum lucidum, dark red; keratin layer may be light blue or light pink or colourless depending upon the tissue and the degree of decolorization. Inner root sheath of the hair, deep blue. Melanin granules, green to black. Other pig- ments, unstained. Bacteria and mycelia, deep blue. Demodex folliculorum in hair follicles, brown with blue granulations. 178 SECTION TWO ORCEIN - PICRO FUCHSIN (Van Gieson) For elastic and collagen fibres Solutions required: A. Orcein (Unna) i% in 80% alcohol 100 ml. Hydrochloric acid, cone. . . i ml. B. Picro-fuchsin (Van Gieson). Technique: 1. Sections are mounted on slides and brought down to 70% alcohol in the usual manner. If tissues have been fixed in a fluid containing mercury, the mercurial precipitate is removed by the standard technique. 2. Immerse in orcein solution (formula as above) for half an hour or longer if necessary ; then rinse in acid alcohol. 3. Rinse in 70% alcohol; then in water. 4. Stain with picro fuchsin (Van Gieson) for three to five min- utes. 5. Rinse rapidly (not more than a few seconds) in water. 6. Dehydrate rapidly; clear, then mount. Results: Collagen fibres are stained red ; elastic fibres, brown ; erythro- cytes, epithelia, muscle, etc., yellow. OSMIC Acm A rapid technique for staining fat in frozen sections Solutions required: A. Osmic acid 1% aqueous. Note: Store in an amber bottle. B. Eosin, yellowish, 1% aqueous. Technique: I. Tissues are fixed as follows : Place 22 5 ml. distilled water in a 50-ml. beaker and heat to about 90° C; then add 2-5 ml. formalin; raise to boiling point; 179 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES drop in a thin piece of the tissue ; then place the beaker in an oven at 60 to 65° C. for ten minutes. 2. Cut frozen sections lo// thick, and place them in another 50-ml. beaker. 3 . Boil Solution A in a large test-tube and pour onto the sections, then transfer to an oven at 60° C. for five minutes. 4. Wash sections in a dish of cold tap water after pouring back the osmic acid, which may be used again, into the stock bottle. 5. Counterstain in Solution B for one minute. 6. Wash quickly in tap water ; transfer sections to slides ; drain and mount in glycerine jelly, previously melted in an oven or on a water bath. Results: Fat globules, black or greyish black against a red background. Caution : Osmic acid vapour is injurious to the eyes. PASINI'S STAIN (Improved) For dififerentiation of connective tissue Solutions required: A. Iron alum 2-5% aqueous. B. PasinVs stain: Unna's aniline blue-orcein . . 10 ml. Eosin bluish 2% in 50% alcohol . . 12 ml. Acid fuchsin . . . . . . 0-3 gm. Neutral glycerine . . . . • • 5 ml- Technique: 1. Tissues should be fixed in Heidenhain's susa mixture and embedded in L.V.N. Sections are cut 3/^ in thickness. After removal of mercuric precipitate in the usual manner sec- tions are mordanted in Solution A for twenty-four hours. 2. Transfer to Solution B for three to ten minutes. 3. Transfer to 95% alcohol and agitate for about one minute or until the colour ceases to come out in clouds. 4. Immerse in absolute alcohol for one minute ; then blot, clear and mount. 180 SECTION TWO Results: Collagen fibres, deep blue. Cytoplasm, red. Epithelial cells, centroiles, basal bodies, nuclear structure, brilliant red. Erythro- cytes, yellowish red. Connective tissue, blue. Secretory bodies, varying according to their nature. Slime of goblet cells, azure blue. Connective tissue, wandering cells, smooth and striated muscle, well defined. PERIODIC ACID (HOTCHKISS) - CELESTIN BLUE For human and animal pituitary glands, demonstrating both muco-protein precursors of the gonadotrophins Compared with other methods, finer differences in the cytology of the cyanophils can be appreciated. Cell counts can readily be carried out, and the counts are more accurate, giving more definite and clearer results than those obtained by older methods : for instance cells appearing by Mallory and other histological methods, to be chromophobes, are found to belong to cyanophil series. The method has been applied with good cytological results to the hypophysis of sheep and goats. Solutions required: A. Helly Fixative containing 5% of neutralized formalin. B. Lugol's iodine. C. Sodium thiosulphate 5% aqueous. D. Hotchkiss' periodic Fuchsin. E. Hotchkiss' reducing rinse {Solution C as page 184). F. Feulgen's Fuchsin. G. Celestin blue {Lendrum and Mcfarlane) Celestin blue . . . . . . 0-25 gm. Iron alum . . . . . . . . 2*5 gm. Distilled water . . . . . . 50 ml. Glycerine . . . . . . • • 7 n^» 181 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Dissolve the alum in the water: then add the celestin blue, and boil for three minutes. Cool and filter ; then add the glycerine. H. Toluidine blue 0*5% aqueous. I. Orange G . . . . . . . . 2 gm. Phosphotungstic acid 5% aqueous. . 100 ml. Allow to stand for 48 hours ; filter before use. Technique: 1. Human and larger animal glands are bisected in the hori- zontal plane with a sharp knife, and the two halves are fixed and embedded separately. Rat hypophyses are fixed in situ after re- moval of the brain and overlaying meninges, by immersing the whole base of the skull in a beaker of the fixative. 2. Fix in Helly for eighteen hours (man, goat, sheep), or two to four hours (rat). 3. Wash from six to twelve hours in running water. 4. Dehydrate, clear and embed in paraffin wax. 5. Cut sections 4-5/x (rat), or 5-5jLt (man). 6. Fix sections to slides; remove paraffin wax with xylol and pass through the usual descending grades of alcohol to distilled water. 7. Immerse in solution B for three minutes. 8. Immerse in solution C (sodium thiosulphate) for three minutes by which time the natural colour of the sections should have been restored and mercurial precipitate removed from the fixative. 9. Wash well with water. 10. Rinse with 70% alcohol. 11. Immerse in periodic acid (solution D) for five minutes. 12. Wash with 70% alcohol. 13. Immerse in Hotchkiss' reducing rinse (solution E) for one minute. 14. Wash with 70% alcohol. 15. Immerse in Feulgen's Fuchsin for ten to thirty minutes. 182 SECTION TWO 1 6. Wash in running water for ten to thirty minutes. 17. Stain in the celestin blue solution one half to three minutes. 18. Rinse in water. 19. Stain in the toluidine blue (solution H) for one half to three minutes. 20. Wash in running water for five minutes. 21. Stain in the phosphomolybdic orange G (solution I) for ten seconds. 22. Wash in water for five to thirty seconds, until a yellow tinge is just visible to the naked eye. 23. Dehydrate through the usual graded alcohols. 24. Clear in xylol and mount. Note : In place of Toluidine blue, Iron Haematoxylin may be used at step 19 in which case it will be necessary to differentiate quickly before washing in running water {step 20). Results : Beta granules in the cyanophils and a number of granules and vesicles in cells which stain as chromophobes by other methods, are magenta to deep red: the colloid is magenta. Alpha granules of acidophils, orange yellow, and erythrocytes are a shade more yellow. Nuclei: are blue-black. Reference: Pearse, Anthony G. Everson (1950), Stain Tech., 25, 95-102. PERIODIC ACID - FEULGEN FUCHSIN (Hotchkiss) For staining polysaccharide structures in fixed animal or plant tissue preparations Solutions required: A. Periodic acid Distilled water Sodium acetate M/5 B. Distilled water Periodic acid 183 0*4 gm. 45 nfil- 5 ml. 10 ml. 0-4 gm. MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Sodium acetate M/5 . . • • 5 ml. Absolute alcohol . . . . 35 ml. Note: This solution, which deteriorates after a few days, should be kept in an amber bottle. C. Reducing rinse: Potassium iodide . . . . i gm. Sodium thiosulphate . . . . i gm. Distilled water . . . . . . 20 ml. Dissolve ; then add with stirring : Absolute alcohol . . . . 30 ml. Hydrochloric acid 2N . . . . 0-5 ml. Note: A precipitate of sulphur is slowly formed and this may be allowed to settle out, or the solution may be used immediately. The solution loses its acid reaction on keeping for some time, and it should be tested with litmus paper ; if the reaction is no longer acid a few drops of N/2 hydrochloric acid should be added until an acid reaction is obtained. D. Feulgen's fuchsin. E. Sulphite wash water: Distilled water . . . . . . 45-5 ml. Hydrochloric acid, pure, cone. . . 0-5 ml. Potassium metabisulphite . . 0-2 gm. Technique: Any fixative may be used but for glycogen or other easily soluble polysaccharides, alcohol is the best fixative as it does not dissolve such substances, and Solution B should be used in place of Solu- tion A. 1. The sections or smears are brought down to alcohol; then immersed in Solution A for five minutes. 2. Pour off solution; flood with 70% alcohol. 3. Immerse for five minutes in Solution C. 4. Flood with 70% alcohol; pour off; then immerse in Solu- tion D for fifteen to forty-five minutes. 5. Wash two or three times in Solution E. 184 SECTION TWO 6. Wash thoroughly in distilled water. 7. Dehydrate ; clear and mount in the usual manner. Results: The following are stained (red) intensely by this reaction: muscle glycogen, liver, hyaluronic acid, gastric mucin, umbilical cord polysaccharide, chitin, crude serum albumin, crude casein, pneumococcus type III polysaccharide, Friedlander type B poly- saccharide, algin, lemon pectin, gum arable, gum tragacanth, gly- cerine, serine, dihydroxyacetone, ribose, arabinose, a-glycero- phosphate, mannitol, tartaric acid, gluconic acid ; while the follow- ing take up the stain with moderate intensity : starch, glucuronic acid, pneumococcus type I polysaccharide, pneumococcus type II polysaccharide; and the following are weakly stained: cellu- lose, crystalline serum albumin, crystalline tgg albumen, glucose, glucosamine, glucose-1-phosphate, galactose, maltose, saccharose, xanthosine, adenosine, muscle adenylic acid and phlorizin. Tryptophane is coloured brown, and the following do not take up the stain: ribonucleic acid, desoxyribonucleic acid, cellobiose, inositol, malic acid and tyrocidine. Notes: {a) Plant tissues are brilliantly stained, in general revealing cellulose or cellulose-like walls of the individual cells and stored carbohydrate such as starch granules, particularly in the region of the chloroblasts if these are present. The cell walls of freshly cut potato reveal every fold, wrinkle or tear. (b) Accumulations of polysaccharides are less common in animal preparations, but mucin, because of its polysaccharide content, is strongly stained. (c) If it is desired to demonstrate glycogen^ Solution B should be used in place of Solution A. From Archives of Biochemistry, Vol. 16, No. i, pages 131-142 (R. D. Hotch- kiss), by courtesy of Academic Press, Inc., New York, 10, U.S.A. PHLOXIN - HAEMATOXYLIN For hyaline, in animal tissues Solutions required: A. Ehrlich haematoxylin solution. B. Phloxin 0.5% in 25% alcohol. 185 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES C. Lithium carbonate o-i% aqueous. Technique: 1. Stain in Ehrlich haematoxylin for five minutes. 2. Wash in water; then stain one half to one minute in the phloxine solution. 3. Wash in tap water; then decolorize in the lithium carbon- ate solution. 4. Wash in tap water; then dehydrate in the usual manner and mount in balsam. Results: Fresh hyaline appears as red droplets and threads ; while older hyaline is pink to colourless ; nuclei, blue. PHLOXIN - METHYLENE BLUE Rapid smear technique for Negri bodies in brain tissue (J. R. Dawson's method) Solutions required: A. Phloxin 2% aqueous. B. Methylene Blue (Loeffler). Technique: 1. The brain to be examined should be removed as quickly as possible; then small segments, 3 to 4 mm. thick, are cut from Ammon's horn perpendicular to its long axis and placed in a Petri dish. Cut away adjacent tissue, leaving only the horn. 2. Place a segment, cut surface downwards, on the small end of a new one-inch cork; then with a matchstick, wipe peripheral tissue downward and outward, so that the segment is more firmly attached to the cork and the grey matter containing the pyramidal cells bulges upwards. Press this gently against a scrupulously clean slide, and make a smear by repeating this process along the whole length of the slide. This operation should be carried out rapidly before the tissue commences to dry. 186 SECTION TWO 3. Fix immediately by immersing in pure methyl alcohol for five to ten minutes. 4. Rinse in running water; then stain in the phloxin solution from two to five minutes. 5. Wash in running water; then stain in methylene blue (Loeffler) for ten to thirty seconds. 6. Decolorize in 80% alcohol; then dehydrate in 95% alcohol and two changes of absolute alcohol. 7. Clear in xylol and mount in balsam. Note: The slides should be handled with forceps throughout to prevent the preparation being spoiled by coming into contact with the fingers. Results: Pyramidal cells, blue; Negri bodies, bright red to reddish brown. PHLOXIN - METHYLENE BLUE - AZUR B For normal and pathological animal tissues This is a rapid modification of Mallory's original method re- ducing the staining time from one hour or more to one to two minutes, and permitting good staining with formalin fixed tissues which is not possible with Mallory's original method which was designed for Zenker-fixed material. Colophony differentiation is obviated, and Phloxin is not washed out as in the original Mallory technique. Solutions required: A. Phloxin . . . . . . . . 0-5 gm. Acetic acid 0-2% aqueous . . . . 100 ml. Filter before use. B. Methylene blue . . . . . . 0-25 gm. Azur B . . . . . . . . 0-25 gm. Borax.. .. .. ., .. 0-25 gm. Distilled water . . . . . . 100 ml. C. Acetic acid 0-2% aqueous. 187 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1 . Fix sections to slides : remove paraffin and pass through the usual descending grades of alcohol to distilled water. 2. Immerse in solution A for one to two minutes. 3. Rinse well in water. 4. Stain for one half to one minute in solution B. 5. Destain partially in solution C. 6. Differentiate in three washes of 95% alcohol. 7. Dehydrate with two changes of absolute alcohol. 8. Clear in two changes of xylol. 9. Mount in synthetic medium (Permount, D.P.X. or Clear- mount, etc.) Results: Nuclei and bacteria are stained blue with some metachromasia. Collagen and other tissue elements are bright pink to red. Erythro- cytes, bright scarlet. Reference: Thomas, John T. (1953), Stain Tech., 28, no. 6. 311 — 312. PHLOXIN - TARTRAZINE (A. C. Lendrum's technique) A general histological stain and for the demonstration of inclusion bodies Note: This technique, in which use is made of a stable phloxine solution, and which affords the advantage of brilliant demon- stration of certain inclusion bodies, is superior to Masson*s erythrosin-saffron, which is less readily prepared and which deteriorates fairly rapidly. Solutions required: A. Haemalum (Mayer). B. Calcium chloride 0-5% aqueous. 100 ml. Phloxine . . . . . . . . 0-5 gm. C. Tartrazine, saturated in cellosolve. 188 SECTION TWO Technique: 1. Stain for five to ten minutes with the haemalum, examining under the microscope at intervals, until the desired depth of staining has been attained. 2. Wash and blue in tap water or in saturated aqueous lithium carbonate. 3. Stain in the phloxine solution for half an hour. 4. Rinse quickly in water. 5. Drain off excess water and replace with tartrazine solution (Solution C, as above), using a dropping bottle to control the differentiation. Note: The tartrazine replaces the phloxine from collagen. As tartrazine is readily soluble in water, slight overstaining is re- commended before dehydration. 6. Rinse in 60% alcohol followed by 95% alcohol. 7. Dehydrate with absolute alcohol. 8. Clear in xylol and mount. Results: Kurloff bodies in guinea pig's lung are well shown. Inclusion bodies of a number of virus-containing tissues show retention of phloxine. Note: Fixatives containing mercuric chloride give the best results. Abstract J'. Path. & Bad., 59, 399-404, 1947, Lendrum, A. C. PICRO - NIGROSIN For eleidin and keratin Solutions required: A. Picric acid, saturated, aqueous. B. Nigrosin 1% aqueous. C. Terpineol . . . . . . . . i volume Origanum oil . . . . . . i volume 189 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Tissues should be fixed in io% formalin, and frozen sections employed. 2. Sections are stained for five minutes in the picric acid solu- tion. 3. Rinse in distilled water. 4. Stain in the nigrosin solution for one minute. 5. Wash in distilled water. 6. Rinse in 95% alcohol. 7. Clear in Solution C. 8. Mount in balsam or in Cristalite. Results: Eleidin, blue-black. Keratin, bright yellow. PROTARGOL - GALLOCYANIN (Foley) For nerve fibres, sheaths and cells Solutions: A. Protargol . . . . . . . . 1% aqueous. (Prepared by sprinkling the protargol powder on the surface of the water and leaving it to dissolve.) B. Protargol 1% aqueous . . . • 50 ml. Alcohol 95% . . . . . . 50 ml. Pyridine pure . . . . . . 0-5 ml. Note: The quantity of pyridine may be varied between o-i ml. and 2 ml. The higher concentrations facilitate the staining of thin fibres, whereas cell bodies and dentrites are better demonstrated with the lower proportions of pyridine. C. Boric acid . . .. .. .. 1-4 gm. Sodium sulphite anhydrous . . 2 gm. Hydroquinone .. .. .. 0-3 gm. Acetone . . . . . . • • 15 rnl. Distilled water . . . . • . 85 ml. Dissolve each reagent in the above order in the water adding the next only after the previous one has been dissolved entirely. 190 . . I ml. . . 0-5 gm . . 2 gm. . . 8 ml. . . 91 ml. SECTION TWO D. Brown gold chloride . . . . 0-2 gm. Distilled water . . . . . . 100 ml. Glacial acetic acid . . . . i ml. Note: Solutions A, B, C and D must be stored in dark bottles. E. Oxalic acid 2% aqueous. F. Sodium thiosulphite 5% aqueous. G. Gallocyanin solution (Einarson). H. Phosphotungstic acid 5% aqueous. I. Aniline Blue 1% aqueous Fast Green FCF . . Orange G Glacial acetic acid Distilled water Note: This solution should be diluted 2 : 3 with distilled water before use. Technique: Note: Metallic instruments must not be used in the following procedures which should be carried out in a darkened room. Sections should be stained in black embryo dishes or in tubes covered all round with thick brown or black paper. Fix in 10% formalin and embed in Celloidin or L.V.N. Sec- tions are cut 15 to 25^ in thickness. 1. Immerse for twenty-four hours in ammoniated alcohol (cone, ammonia i ml., alcohol 50% 99 ml.). 2. Drain well and transfer to Solution A for six to eight hours at 37° C. 3. Drain well and transfer to Solution B in another staining dish, for twenty-four to forty-eight hours at 37° C. 4. Rinse for about five to ten seconds in 50% alcohol; then reduce with Solution C for about ten minutes. 5. Wash in several changes of distilled water; then tone in Solution D for ten minutes. 6. Wash in several changes of distilled water ; then transfer for one to three minutes in Solution E, afterwards rinsing in distilled water. 191 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 7. Immerse for three to five minutes in Solution F ; then wash thoroughly in distilled water. 8. Counterstain overnight in Solution G. 9. Wash thoroughly in distilled water then immerse in Solution H for thirty minutes. 10. Without washing transfer section to diluted Solution I for one hour; then wash with 70% alcohol and differentiate the counterstain in 95% alcohol. 11. Transfer to normal butyl alcohol; clear in cedarwood oil and mount. Results: Nerve fibres and neurofibrils, blue-black. Nissl bodies, pale blue. Nuclei, blue-black with silver and gold if a higher percen- tage of pyridine was used in solution. Myelin sheaths, bright yellow. Connective tissues, various shades of blue and green. PURPURIN For calcium deposits in pathological tissues Solutions required: A. Purpurin, saturated in absolute alcohol (about 0*7%). B. Sodium chloride, reagent grade, 0-75% aqueous. Technique: 1. Fix material in 90% alcohol. 2. Dehydrate; clear; embed in paraffin wax. 3. Fix sections to slides; de-wax and pass through the usual descending grades of alcohol to distilled water. 4. Stain in the purpurin solution for about ten minutes. 5. Immerse in sodium chloride solution for about five minutes. 6. Rinse with 70% alcohol until the stain ceases to come away in clouds. 7. Rinse with 90% alcohol and dehydrate with absolute alcohol. 8. Clear in xylol and mount in balsam. Results: Calcium deposits are stained red. 192 SECTION TWO QUINCKE REACTION For haemosiderin Solutions required: A. Ammonium sulphide solution, concentrated . . . . . . i volume Absolute alcohol . . • • 3 volumes B. Basic fuchsin 0-5% in 50% alcohol. Technique: 1. Tissues are fixed in neutral formalin 10%, or in absolute alcohol, and embedded in paraffin wax or Celloidin in the usual manner. 2. Bring sections down to distilled water, then immerse them from two to forty-eight hours in the ammonium sulphide solution. 3. Rinse thoroughly in distilled water. 4. Counterstain in the basic fuchsin solution for five to twenty minutes. 5. Wash in water; drain well; rinse in 80% alcohol. 6. Differentiate and dehydrate in absolute alcohol; clear in xylol and mount in Canada balsam or in Cristalite. Results: Haemosiderin, dark brown to black. RHODAMINE B - ANILINE METHYLENE BLUE For splenic and lymphoid tissues Solutions required: A. Methylene Blue, 2% alcoholic Aniline water . . ^ . . Distilled water B. Rhodamine B 1% aqueous Distilled water C. Solution A (above) Solution B (above) 193 10 ml. 15 ml. 30 ml. 2-5 ml. 47-5 ml. 3 volumes 7 volumes MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: Tissues are fixed in Zenker-Formol and embedded in paraffin wax. 1. After removal of mercurial precipitate by treatment with iodine in the usual manner {see page 28) sections are stained two to three hours in Solution C; then washed rapidly with absolute alcohol. 2. Clear in xylol ; mount. Results: Basophile protoplasm, blue. Chromatin, violet blue. Nucleoli, red. Connective tissue, faintly stained yellowish red. Muscle, yellowish red. Erythrocytes, bright red. Acidophile granules of leucocytes, bright red. Hyalin and granules of Russell, bright red. Nuclei of the small lymphocytes are faintly stained violet. SAFFRON For connective tissue Note: Saffron is the dried stigmata of crocus sativus, and should not be confused with safranin, which is an aniline dye. Solution required: A. Saffron . . . . . . . . 2 gm. Distilled water . . . . . . 100 ml. Boil gently for an hour ; allow to cool ; then filter ; add I ml. of 40% formaldehyde and i ml. of 5% tannic acid to the filtrate. Note: Saffron solution deteriorates after a few weeks, and it is best to prepare the solution in small quantities, as required. B. Delafield or Ehrlich haematoxylin. C. Erythrosin, 1% aqueous. Technique: 1. Fix small pieces of tissue in Bouin, Zenker-formaldehyde or in mercuric-formaldehyde. 2. Wash; dehydrate; embed. 194 SECTION TWO 3. Sections are stained for five to ten minutes with Delafield or Ehrlich haematoxylin ; rinse in water. 4. Blue in tap water in the usual manner or in 1% sodium phos- phate (Na2HP04). 5. Stain for two to five minutes in 1% aqueous erythrosin. 6. Rinse quickly with water. 7. Differentiate with 70% alcohol for a few seconds, controlling under the microscope, until the collagen fibres are nearly colour- less. 8. Rinse in water; stain for five minutes in saffron solution prepared as above ; rinse with water. 9. Wash rapidly first with 70% alcohol then with absolute alcohol ; clear in xylol and mount. Results: Nuclei are stained blue. Cytoplasm, varying shades of red. Muscle, pink. Elastic fibres, pink. Collagen, yellow. Improved differentiation of most cells and tissues is obtained by employing this method in place of haematoxylin and eosin. SAFRANIN - CRYSTAL VIOLET - FAST GREEN - ORANGE 2 (S. S. Kalter's " Quadruple Stain for Animal Tissues ".) Abstract from jf. Lab. & Clin. Med., 28, 995-7, 1943. This technique, which is a development of Flemming's triple stain, is particularly useful for histology students Solution required: A. Safranin, O . . . . . . 0-2 gm. Formalin (40% formaldehyde) . . 4 ml. Sodium acetate . . . . . . 0-5 gm. Alcohol 50% . . . . . . 100 ml. B. Crystal violet 0-5% aqueous. C. Fast green F.C.F. - Orange G saturated in clove oil D. Orange 2 saturated in clove oil. 195 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Any fixative may be employed, but if Bouin is chosen, remove picric acid with a few drops of saturated lithium carbonate in the 70% alcohol of dehydration series. Should a fixative containing mercuric chloride be chosen, then it will, of course, be necessary to remove mercuric precipitate in the usual way. 2. After fixation, wash, dehydrate, clear and embed in paraffin wax. 3. Cut sections no thicker than 7-5// and fix to slides, avoiding the use of glycerine albumin, which will cloud the stain. 4. De-wax with xylol ; then pass through absolute, 90% and 70% alcohol to water. 5. Stain in the safranin solution for twenty-four hours. 6. Rinse with water. 7. Stain with the crystal violet for one to two minutes. 8. Wash with water. 9. Immerse in 50% alcohol for two minutes. 10. Immerse in 95% alcohol for two minutes. 11. Immerse in the fast green-orange 2 (Solution C) for five minutes. 12. Differentiate, examining under the microscope, until the connective tissue is stained to the desired depth of green. 13. Immerse in clove oil for ten minutes. 14. Transfer to Orange 2 in clove oil for ten minutes. 15. Differentiate, examining under the microscope at intervals, until the desired depth of staining has been achieved. 16. Immerse for ten minutes each in two changes of xylol. 17. Mount in balsam. Results: Nuclei, red. Nucleoli, purple or purpHsh red. Nuclear mem- brane, dark red. Cellular cytoplasm, pink to red, except in Henle's loop (light green). Connective tissue, green. Elastic fibres, yellow. Fibroblasts, green with purple nuclei. Muscle, reddish brown. Erythrocytes, orange. Polymorphonuclears show purple nuclei. 196 SECTION TWO SAFRANIN - WATER BLUE (Unna) For collagen fibres Solutions required: A. Safranin O i% aqueous. B. Water Blue i% aqueous. . . . lo ml. Tannic acid 33% aqueous . . 10 ml. This solution must be freshly prepared. Technique: 1. Tissues should be fixed in 1% aqueous picric acid or in absolute alcohol, and Celloidin sections should be employed. 2. Stain for ten minutes in the safranin solution; then wash thoroughly in water. 3. Stain for ten to fifteen minutes in Solution B. 4. Wash thoroughly in distilled water. 5. Clear in Bergamot oil; then mount. Results: Collagen fibres are stained blue, while nuclei are red SCARLET R - ETHYLENE GLYCOL An improved technique for staining fat, etc, in animal tissues, its chief advantages being: (a) Excellent differentiation without loss of stain out of the lipid particles, {b) A stable solution which does not dissolve lipid materials, {c) Sections are not shrunken but remain pliable, {d) More intense staining of fat. Solutions required: A. Ethylene glycol, pure, anhydrous. B. Scarlet R . . . . ^ . . . . i gm. Ethylene Glycol, pure, anhydrous. . 100 ml. Heat the ethylene glycol to 100-110° C. on a hot plate or in an oven, or over the bunsen flame, taking care that it does not catch fire; then add the stain and stir until all or most of it is dissolved. Filter when cold. 197 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES . C. Ethylene glycol, pure, anhydrous . . 85 ml. Distilled water . . . . . . 15 ml. D. Ehrlich or Delafield haematoxylin. Technique: 1. Fix material in 10% formalin and cut frozen sections. 2. Wash sections in water for two minutes or longer to remove the formalin. 3. Dehydrate the sections by agitating gently in pure anhydrous ethylene glycol for three to five minutes. 4. Immerse the sections in the stain (solution B) for two to three minutes, with gentle agitation. 5. Differentiate by agitating gently in 85% ethylene glycol (solution C) from one to ten minutes, controlling by examination under the microscope while the preparation is still wet. 6. Transfer to distilled water for three to five minutes. 7. Counterstain with Ehrlich or Delafield haematoxylin. 8. Wash well in tap water. 9. Mount in glycerine jelly. Results: Nuclei, blue. Fat, orange to red. Cholesterol, red. Normal myelin, unstained. Fatty acids, unstained. SCARLET R For staining fat, etc. in animal tissues Solutions required: A. Scarlet Red, saturated in equal volumes of acetone and 70% alcohol. B. Ehrlich or Delafield haematoxylin. Technique: I. Tissues are fixed in formalin and frozen sections are em- ployed. 198 SECTION TWO 2. Sections are immersed for a second in 70% alcohol; then stained for two to five minutes, in the Scarlet R solution. 3. Wash quickly in 70 % alcohol, and transfer to distilled water. 4. Counterstain with Ehrlich or Delafield haematoxylin. 5. Wash well in tap water; mount in glycerine or glycerine jelly. Results: Nuclei, blue. Fat, orange to red. Cholesterol, red. Normal myelin, unstained. Fatty acids, unstained. SILVER CARBONATE - ORCEIN - ANILINE BLUE - FAST GREEN For demonstrating reticulin, elastin and collagen in the same tissue sections Solutmis required: A. Celloidin 0-5% in equal vols, of Ether and Absolute Alcohol. B. Pot. Perman. 0-25% aqueous. C. Oxalic Acid 5% aqueous. D. Silver carbonate^ Hortega {Foot's modification) Silver nitrate 10% aqueous . . 10 ml. Lithium carbonate, saturated aqueous . . . . . . . . 10 ml. Shake well; then allow to stand for ten minutes or so in a 25 ml. measuring cylinder. Pour off the supernatent fluid, then transfer the precipitate to a 100 ml. measuring cylinder, and add about 75 ml. distilled water, shake well ; allow to settle ; then pour off the fluid and add a second lot of distilled water. This process should be repeated three or four times. Finally add 25 ml. of distilled water to the precipitate and add 28% ammonia solution drop by drop until the precipitate is almost dissolved. Make up to 100 ml. with 90% alcohol. Filter and warm to 50° C. for 15 minutes before using. p 199 ' MEDICAL AND BIOLOGICAL STAINING TECHNIQUES E. Neutral Formalin 40% . . . . 20 ml. Distilled water . . . . . . 80 ml. Buffer to pH 7-0. F. Gold chloride 0*2% aqueous. G. Sodium Thiosulphate 5%. H. Orcein 1% in 70% alcohol . . 100 ml. Hydrochloric acid, cone. . . . . i ml. Picro Aniline Blue I. Aniline blue aqueous . . . . o*i gm. Picric acid, saturated aqueous . . 100 ml. or: J. Picric acid, saturated aqueous . . 100 ml. Fast Green FCF . . . . . . 0*2 gm. Technique: 1. Material should be fixed in 10% formalin and embedded in paraffin wax by the standard technique. 2. Sections 4 to 5/i, in thickness are fixed to slides, dewaxed and immersed in xylol for five minutes. 3. Wash and immerse in absolute alcohol for two minutes. 4. Immerse in solution A for five minutes (Celloidin). 5. Drain slides for one minute. 6. Immerse in 80% alcohol for five minutes. 7. Rinse in water. 8. Immerse in solution B (Pot. perman.) for five minutes. 9. Rinse in water. 10. Immerse in solution C (oxalic acid) for five minutes. 11. Rinse in tap water. 12. Wash in distilled water. 13. Immerse in the silver carbonate solution in an oven at 50° C. for ten to fifteen minutes. (Soln. D). 14. Rinse in distilled water. 15. Immerse in solution E (20% neutral formalin) for five minutes. 200 SECTION TWO 1 6. Rinse in tap water. 17. Tone in solution F (Gold chloride) for five minutes. 18. Rinse in tap water. 19. Immerse in solution G for two minutes. 20. Rinse in tap water. 21. Stain in solution H (Orcein) for ten to fifteen minutes in an oven at 37° C. or for one hour at room temperature. 22. Rinse in 70% alcohol, followed by tap water. 23. Stain in solution I (Aniline blue) for twenty to forty seconds or solution J (Fast green) for ten to twenty seconds. 24. Wash with 95% alcohol for six to eight seconds. 25. Rinse briefly with absolute alcohol. 26. Rinse with a mixture consisting of equal volumes of xylol and absolute alcohol until clear. 27. Rinse with several changes of xylol, and mount. Results: > With Picro aniline blue Elastic fibres reddish brown Collagen blue Reticulum black Muscle blue-green Nuclei tan to brown Erythrocytes yellowish tan Cytoplasm pale blue With Picro fast green Elastic fibres orange to reddish brown Collagen blue-green Reticulum black Muscle green Nuclei tan to brown Erythrocytes yellow to orange Cytoplasm light green Reference: Lewis, Ann L., and Jones, Russell S., (1951), Stain Tech., 26, 85-7. 201 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES SILVER NITRATE - GOLD CHLORIDE - PARACARMINE (Da Fano) For Golgi apparatus A. Cobalt nitrate i% aqueous . . lOO ml. Formalin . . . . . . • • 15 ml- B. Silver nitrate 1*5% aqueous. (N.B. : This should be stored in an amber or blue glass bottle.) C. Cajal's Reducer. Hydroquinone 2% aqueous . . 100 ml. Neutral formalin . . . . . . 15 ml. Sodium sulphite anhydrous . . 0-5 gm. N.B.: This solution should be freshly prepared. D. Gold chloride 0-2% aqueous. E. Sodium thiosulphate 5% aqueous. F. Paracarmine (Mayer). * Technique: 1. Pieces of tissue no thicker than 3 mm. are fixed from two to eighteen hours in the cobalt nitrate formalin solution, according to the size and nature of the material. 2. Wash the tissue quickly in a large volume of distilled water- 3. Immerse in the silver nitrate solution in the dark for thirty- six to forty-eight hours. 4. Wash quickly in a large volume of distilled water. 5. Trim the tissue to a thickness not exceeding 2 mm. 6. Immerse in Solution C (Cajal's Reducer) for two to twenty- four hours in the dark. Note: For most soft tissues about four hours will suffice. 7. Wash in several changes of distilled water. 8. Dehydrate, clear and embed in paraffin wax in the usual way. 9. Cut sections up to Sju in thickness. 202 SECTION TWO 10. Fix sections to slides ; de-wax and pass through descending grades of alcohol down to distilled water. 11. Tone sections on slides by immersing in the gold chloride solution for five to ten minutes. 12. Wash quickly in distilled water. 13. Fix in 5% sodium thiosulphate (Solution E) for ten to fifteen minutes. 14. Wash thoroughly in distilled water. 15. Counterstain with paracarmine for about ten minutes. 16. Rinse with 90% alcohol, followed by absolute alcohol. 17. Clear in xylol and mount. Results: Golgi apparatus stained black while cells are pink or red. SILVER NITRATE - HYDROQUINONE For the detection of gold in filxed tissues of experimental animals Notes: (a) The following technique must be carried out in the dark- room. (b) Fixatives containing metals must be avoided. {c) The use of metal instruments in handling the sections must be avoided. Solutions required: A. Gum acacia 10% aqueous, filitered 100 ml. Silver nitrate, A.R. grade . . 2 gm. Note: This solution should be prepared immedi- ately before use. B. Gum acacia 10% aqueous, filtered 100 ml. Hydroquinone . . . . . . i gm. Note: This solution should be prepared the day before it is required for use. 203 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES C. Solution A . . . . . . lo ml. Solution B . . . . . , 10 ml. Citric acid 5% . . . . . . 0-5 ml. Note: This solution should be prepared only when required for immediate use. D. Sodium thiosulphate 5% aqueous. Technique: 1. Small pieces of tissue are fixed in 20% formalin, and frozen sections are employed. 2. Rinse thoroughly in distilled water. 3. Immerse sections in Solution C and leave therein for five to ten minutes. 4. Plunge sections directly into the sodium thiosulphate solu- tion, without prior washing, and leave therein for five minutes. 5. Wash thoroughly in several changes of distilled water. 6. Mount in Aquamount or in Farrant's medium. Results: The presence of gold is indicated by a black deposit in the cells. SUDAN BLACK For lipids (especially those that are not well coloured by Sudan III or IV) (J. R. Baker's technique) Solutions required: A. Formaldehyde-saline. Formalin (Formaldehyde 40%) . . 10 ml. Sodium chloride 10% aqueous . . 7 ml. Distilled water . . . . . . 83 ml. Note: Keep a few pieces of marble chips in the solution to maintain neutrality. B. Formalin (Formaldehyde 40%) neutral. Note : Keep a few pieces of marble chips in the bottle, 204 SECTION TWO C. Potassium dichromate 2-5% aqueous . . . . . . . . 88 ml. Sodium chloride 10% aqueous . . 7 ml. Note: Keep a few pieces of marble chips in the bottle. D. Dichromate-formaldehyde. Solution B I volume. Solution C 19 volumes. E. Potassium dichromate 5% aqueous F. Gelatine for embedding. Gelatine powder . . . . . . 25 gm. Water . . . . . . . . 100 ml. Sodium p-hydroxybenzoate . . 0-2 gm. Sprinkle the gelatine on to the water and leave it to soak for an hour, afterwards warming in an incubator maintained at 37° C. until all the gelatine has dissolved, then strain through muslin while still warm. Note: If Sodium p-hydroxybenzoate, which is added to prevent the growth of moulds and bacteria, is not available in the laboratory, then 0-25 to 0*5 gm. of Thymol should be used instead. G. Formalum {for hardening gelatine) Formalin (Formaldehyde 40%) . . 20 ml. Potassium alum 5% aqueous . . 80 ml. Keep marble chips in the bottle. Note: Both gelatine blocks and gelatine sections may be pre- served indefinitely in Formalum, which makes the gelatine very hard, thereby facilitating the cutting of thin sections which are non-sticky. Important: Formalum must not be used in the acid haematein test for phospholipids (Baker 1946) as the alum would react with the haematein. H. Sudan Black ... ... . . 0-5 gm. Alcohol 70% 100 ml. Boil for ten minutes under a reflux condenser ; then cool and filter. I. Car malum {Mayer). 205 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Fix a piece of tissue not more than 3 mm. thick in the for- maldehyde-saline for an hour. 2. Transfer, without washing, to the dichromate formaldehyde (Solution D) and leave for five hours. 3. Transfer, without washing, to 5% aqueous potassium dichromate and leave for about eighteen hours. 4. Leaving the tissue in the same solution, transfer to the paraf- fin oven at 60° C. for twenty-four hours. 5. Wash in running water for six hours. 6. Leave overnight in the melted gelatine in the oven at 37° C. 7. Cool the gelatine, preferably in a refrigerator. 8. Cut out a rectangular block containing the specimen. 9. Immerse the block overnight (or any conveniently longer time) in formalum, placing a marble chip in the capsule or tube. 10. Cut sections 8 to lOju, on the freezing microtome. 11. Transfer a section to 70% alcohol. Note: It is best to transfer sections from fluid to fluid, up to stage 16 in a Royal Worcester Porcelain thimble No. ^.4756, size 2. 12. Transfer to the Sudan black solution, and leave for |~4 minutes. (The best period is usually about 2 J minutes.) 13. Wash in 70% alcohol for five seconds. 14. Wash in 50% alcohol for one minute. 15. Wash in water, sinking the section gently with a camel hair brush if it floats. 16. Transfer to Carmalum for two to three minutes. (The optimum time is usually three minutes.) 17. Rinse in distilled water. 18. Transfer the section to a fairly large dish, or a tongue jar of tap water, and leave for two minutes, or any conveniently longer time. 18. Wash again in another large bowl of water. 19. Transfer to a petri dish of water. 20. Float the section on to a slide. 206 SECTION TWO 21. Blot away excess water but do not allow the section to dry. 22. Mount in Farrant's medium, or in Aquamount. 23. Attach a clip to hold the coverslip to the slide: then leave overnight in the oven to harden the mounting media, before examining the preparation under the oil immersion objective. Note: The slide may be examined after a quarter of an hour, if desired; then returned to the oven to complete the hardening. Results* : Lipids, dark blue or blue-black. Cytoplasm : colourless or pale grey-blue. Chromatin: pink or red. Note: If the results are not good, another section should be tried with variations of the staining times. Never attempt to judge the colouring until the section is mounted and examined under the oil immersion objective. It is recommended that the technique be learned on the intestine of the mouse, as it is scarcely possible to fail with this. Cut out a piece of empty intestine about i cm. long and immerse in for- maldehyde-saline for five minutes, then open it by a longitudinal cut from one end to the other, taking care not to do any unneces- sary damage to the villi. The section should be left only one minute in the Sudan black and two and a half minutes in the carmalum. Reference: Baker, J. R. (1949), Q.jf.M.S., 90, 293-307. * In the original paper the method refers to " Golgi bodies ". Dr. J. R. Baker, to whom I am indebted for permission to include this abstract, informs me (March, 1955) that he regards the method as essentially for those lipids that are not well stained by other Sudan colours. In the original paper it is stated: " Simple Golgi bodies and externa: dark blue or blue-black. Golgi vacuoles: colourless ". The changes are Dr. J. R. Baker's. SUDAN BLACK A specific stain for neutral fats Solutions required: A. Sudan black, saturated in 70% alcohol. B. Carmalum (Mayer). Technique: Tissues are fixed at least three days in 10% formalin; then rinsed thoroughly in distilled water. 207 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 1. Frozen sections are immersed one minute in 50% alcohol; then one minute in 70% alcohol. 2. Stain for fifteen minutes to several hours in the Sudan black. 3. Rinse for a few minutes in 50% alcohol; then in distilled water. 4. Counterstain in carmalum for about three minutes; wash with distilled water ; mount in glycerine jelly. Results: Neutral fat and myelin, blue-black to black; nuclei, red. SUDAN BLACK - ETHYLENE GLYCOL An improved technique for lipid staining, offering the ad- vantage of a stable solution, excellent differentiation v^^ith- out loss of stain out of the lipid particles, and pliable unshrunken sections Solutions required: A. Ethylene glycol, pure, anhydrous. B. Sudan black . . . . . . . . i gm. Ethylene glycol, pure, anhydrous. . 100 ml. Heat the ethylene glycol to 100-110° C. on a hot plate or in an oven, or over a bunsen flame, taking care that it does not catch fire ; then add the stain and stir until all or most of it is dissolved. Filter when cold. C. Ethylene glycol, pure, anhydrous. . 85 ml. Distilled water . . . . . . 15 ml. D. Carmalum (Mayer). Technique : 1. Fix tissues for at least three days in 10% formalin. 2. Wash thoroughly in running water to remove the fixative. 3. Dehydrate the sections by agitating gently in pure anhydrous ethylene glycol for three to five minutes. 4. Immerse the sections in the Sudan black solution from fifteen minutes to one hour, agitating gently at intervals. 5. Differentiate by agitating gently at intervals with 85% ethylene glycol (solution C) from one to ten minutes, controlling under the microscope while the section is still wet. 208 SECTION TWO 6. Transfer the sections to distilled water for three to five minutes. 7. Counterstain with carmalum for about three minutes. 8. Wash with distilled water. 9. Mount in glycerine jelly. Results: Lipid substances or particles are stained blue-black to black. Nuclei, red. Reference: Chiffelle, T. L., and Putt, F. A. (195 1), Stain Tech., 26, no. i, pages 51-6. SUDAN BLUE For demonstrating degenerated myelin Solution required: Sudan blue, saturated in 50% alcohol. Technique: 1. Tissues should be fixed for at least three days in 10% neutral formalin. 2. Frozen sections are soaked one minute in 50% alcohol; then one minute in 70% alcohol ; then stained from fifteen minutes to several hours in a saturated solution of Sudan blue in 50% alcohol. 3. Rinse for a few minutes in 50% alcohol; then in distilled water. 4. Counterstain in carmalum for three to five minutes. 5. Rinse in water; mount in glycerine jelly. Result: Degenerated myelin, blue. SUDAN BROWN For fat and for acute fatty degeneration not shown by Scarlet R Solutions required: A. Stock solution of Sudan brown saturated in isopropyl alcohol. B. Mayer's acid haemalum. 209 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Fix tissues in io% formalin. 2. Frozen sections are stained for five minutes in a diluted solution of Sudan brown prepared by mixing 6 volumes of the stock solution with 4 volumes distilled water. {Note: This diluted solution keeps only for one day, and should, therefore, only be prepared immediately before use.) 3. Float out in tap water. 4. Stain for two to five minutes in Mayer's acid haemalum; then rinse in tap water. 5. Immerse in tap water or in 1% sodium phosphate (Na2HP04) until the section turns blue. 6. Rinse in distilled water. 7. Mount in Apathy's gum syrup. Results: Fat, brown; nuclei, bluish grey; protoplasm, colourless. SUDAN 2 For degenerating and intact myelin and fat Solutions required: A. Haematoxylin 1% in absolute alcohol, at least one to five days old . . . . . . . . 50 ml. 4% iron alum aqueous . . . . 50 ml. This solution should be prepared immediately before use. B. Borax . . . . . . . . i gm. 1 Potassium ferricyanide . . • • 5 gni- Distilled water . . . . . . 100 ml. 210 SECTION TWO C. Iron alum . . . . . . 0-5% aqueous D. Sudan 2, saturated in isopropyl alcohol . . . . . . . . 30 ml. Distilled water . . . . . . 20 ml. Mix well and allow to stand for ten minutes before use. Note: This solution deteriorates within three to four hours. Technique: 1 . Formalin-fixed frozen sections are stained forty minutes with Solution A at 56° C. in a covered dish in an oven. 2. Rinse in water and differentiate for one hour with Solution B. 3. Rinse in distilled water and immerse for ten minutes in Solution C. 4. Stain ten to twenty minutes in Solution D ; then float out in water. 5. Mount in Apathy's gum syrup or in Aquamount. Results: Normal myelin, blue-black; nerve cells, grey; nuclei, deeper grey ; red corpuscles, yellow to black ; fats, orange yellow. THIONIN (Ehrlich) For mucin Solution required: Thionin (Ehrlich), saturated, aqueous . . . . . . 0-5 ml. Distilled water . . . . . . 10 ml. Technique: I. Tissues should be fixed in Zenker's fluid, washed in running water, dehydrated, cleared, embedded and sectioned in the usual manner. 211 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 2. Sections are mounted on slides and the mercuric precipitate from the fixative is removed by the standard technique {see page 28). 3. Bring down to distilled water as usual; then stain from five to fifteen minutes in the thionin solution. 4. Dehydrate rapidly as otherwise the stain will be removed by the alcohol. 5. Clear in xylol and mount. Results: Mucin is metachromatically stained purple; while the basophil granules of the mast cells, Wharton's jelly of the umbilical cord, are purple; and the other tissue constituents are stained in varying shades of blue. THIONIN (Ehrlich) For the differential staining of entamoeba in sections Solutions required: A. Thionin 0-25% aqueous. B. Oxalic acid 2% aqueous. Technique: 1. Pieces of tissue are fixed in absolute alcohol and embedded in Celloidin in the usual manner. 2. Immerse in the thionin solution for three to seven minutes. 3. Differentiate in the oxalic acid solution for thirty to ninety seconds, controlling by examination under the microscope. 4. Rinse in water. 5. Rinse in 70% alcohol. 6. Dehydrate by rinsing in two changes of 95% alcohol. 7. Clear in terpineol and mount. Results: Nuclei of amoebae are stained a rich brown colour, while the nuclei of all other cells are stained blue. 212 SECTION TWO THIONIN (Ehrlich) For nerve cells and fibre tracts Solution required: Stain and fixative combined: Thionin 0-5% in io% formalin. Technique: 1. Tissues are fixed and stained simultaneously by immersing in the above solution from a few days to three months. 2. Wash well in running water. 3. Dehydrate in ascending grades of alcohol in the usual man- ner. 4. Clear ; embed in paraffin wax or in Celloidin. 5. Fix sections to slides and mount in Cristalite. Results: Cell bodies are stained blue, while the fibre tracts are red. THIONIN (Ehrlich) For demonstrating malignant cells in biopsy material Solution required: Thionin 1% aqueous. Technique: 1. Stain frozen sections for ten to sixty seconds in the thionin solution. 2. Rinse in water. 3. Mount in tap water or in Aquamount. Results: Nuclei, blue to purple. Collagen, red. Elastic tissues, green. 213 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES TOLUIDINE BLUE For mucus Solution required: Toluidine Blue 1-5% aqueous. Technique: 1. Formalin-fixed material is embedded in paraffin wax in the usual manner. 2. Bring sections down to distilled water then stain for one or two minutes in the toluidine blue solution. 3. Wash with distilled water; drain well, then plunge the slide into two changes of 95% alcohol. 4. Dehydrate by immersing in two changes of acetone; then clear in xylol and mount. Results: Mucus, reddish violet; nuclei, blue; erythrocytes, yellow to greenish yellow. TRICHROME STAIN (G. Gomori) A. Delafield or Ehrlich Haematoxylin. B. Lithium carbonate 1% aqueous. C. Alcohol 70% 97 ml. HCl, concentrated . . . . • • 3 nil- D. Picric acid 1% in 50% alcohol. E. Phosphotungstic Acid 3%. F. Light green, or Fast green FCF, or Aniline blue . . . . . . . . 0-5 gm. Neoponceau (Michrome) . . . . 1-5 gm. | G. Solution F . . . . . . . . i volume Acetic acid 2% . . . . 3 to 4 volumes 214 SECTION TWO Technique: 1. Fix tissues in Bouin or io% Formalin. 2. Stain sections in the Haematoxylin solution for ten minutes. 3. Blue in the lithium carbonate solution. 4. Differentiate, if necessary, with the HCl alcohol for pre- dominance of the green or blue shades, or in picric alcohol for predominance of red shades, in the final picture. 5. Immerse the preparation in 3% phosphotungstic acid for ten to fifteen minutes. 6. Wash gently under the tap for one minute. 7. Stain in solution G for five to twenty minutes. Note: The time is not critical although results will be slightly different. 8. Rinse in 2% acetic acid. 9. Dehydrate, clear and mount. Results: Nuclei, blue. Cytoplasm, muscle fibres, red cells, etc., in shades of red. Connective tissue green or blue. From personal communications with Professor G. Gomori of the Department of Medicine ^ University of Chicago, U.S. A., to whom I am indebted for permission to include this hitherto unpublished technique. Note: Professor Gomori has used Woodstain scarlet, which is not available under that name in Britain. The British equivalent is Neoponceau and I feel that this synonym is more suitable in this case as the name Woodstain scarlet, indexed in literature on general biology, might suggest a botanical stain for woody tissues. TRICHROME STAIN (Masson), Modified For epithelium, pituitary and thyroid glands, nerve (normal and tumour), etc. Solutions required: A. Regaud's haematoxylin. B. Picric acid saturated in 95% alcohol 20 ml. Alcohol 95% . . . . . . 10 ml. Q 215 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES C . Violamine-acid fuchsin : Violamine, R . . . . . . 0-7 gm. Acid fuchsin . . . . . . 0-3 gm. Glacial acetic acid . . . . i ml. Distilled water . . . . . . 100 ml. D. Acetic fast green Fast green, F C F . . • • 3 gm- Glacial acetic acid . . . . 2 ml. Distilled water . . . . . . 100 ml. Technique : 1 . Tissues are fixed in Bouin, Regaud, Zenker or formalin, and embedded in paraffin wax. 2. Mordant sections on slides with 5% iron alum previously heated to 45° C. 3. Wash with tap water; stain for five minutes in Regaud's haematoxylin ; then rinse with 95% alcohol. 4. Differentiate with picric alcohol; then wash with running tap water. 5. Stain for five minutes in Ponceau-acid fuchsin; then wash with distilled water. 6. Differentiate for five minutes in 1% phosphomolybdic acid; then without rinsing: 7. Flood the slide with solution D and leave for five to ten minutes. 8. Rinse with distilled water; then return to 1% phospho- molybdic acid for five minutes. 9. Leave in 1% acetic acid for five minutes. 10. Dehydrate in 95% alcohol, followed by absolute alcohol; clear in xylol and mount. Results: Nuclei, black. Argentaffin granules, black or red. Cytoplasm, vermilion. Collagen, green. NeurogUa fibrils, vermilion. Mucus, green. Keratin, vermilion. Intercellular fibrils, vermilion. Golgi apparatus, clear. 216 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES UREA SILVER NITRATE For nerve fibres and nerve endings Note: In this technique, nerve fibres and nerve endings of the peripheral and central nervous system are preferentially stained. Applied to paraffin sections on slides, the technique gives rapid and constant results, and eliminates the necessity of gold toning. The following fixatives are recommended: (I). Chloral hydrate . . . . • • 25 gm. Alcohol 50% . . . . . . 100 ml. (II). Formalin, undiluted (i.e. Form- aldehyde 40%). . . . . . . 20 ml. Alcohol 95% 80 ml. (III). Ammonium bromide . . . . 2 gm. Formalin, undiluted (i.e. Form- aldehyde 40% . . . . . . 15 ml. (IV). 95% or Absolute Alcohol (V). Bouin's fluid. Solutions, I, II, III and IV are satisfactory for Central Nervous System and nerve trunks. Solution I has been used with satisfaction for striated muscle tissue. Solution II is suitable for gland and smooth muscle tissue. Solution V for gland and smooth muscle tissues and for embryos. Tissues may also be fixed in 10% formalin with good results, but an excessive staining of connective tissue has been observed when this fixative has been employed. Solutions required: A. Picric acid, saturated aqueous Mercuric cyanide . . B. Silver nitrate 1% aqueous Urea . . Solution A . . C. Hydroquinone Urea . . Distilled water Sodium sulphite, anhydrous 50 ml. 0-5 gm. 100 ml. 25 gm. 3 drops 2gm. 25 gm. 100 ml. 10 gm. 217 SECTION TWO Technique: 1. Fix the material in one of the above fixatives (I, II, III, IV or V) and embed in paraffin wax. 2. Fix sections to sUdes and remove paraffin wax with xylol. 3. Rinse with two changes of absolute alcohol. 4. Wash with 90% followed by 80% alcohol. 5. Immerse slides directly into solution B for one to one and a half hours at 50 to 60° C. in an oven. 6. Rinse quickly in two changes of distilled water. 7. Reduce by immersing in solution C for three minutes at 25 to 30° C, agitating the slides gently for the first two minutes. 8. Wash thoroughly in four or five changes of distilled water. 9. Wash with 50% followed by 70% and 80% alcohols. 10. Examine under the microscope while the preparation is still wet and if it is found that the staining is not complete, repeat step 5 using the original urea-silver nitrate solution and reducing the time to ten to fifteen minutes ; then repeat steps 6, 7, 8 and 9. 11. Rinse with 95% alcohol. 12. Dehydrate with two changes of absolute alcohol. 13. Clear in xylol and mount. Results: Nerve fibres are stained from brown to black, while nerve end- ings are usually black, and nerve cells from yellow to brown. The background is usually yellow, but its appearance depends upon the kind of tissue and the fixative employed. Reference: Ungewitter, L. H. (195 1), Stain Tech., 26, p. 75. VERHOEFF'S STAIN For elastic fibres, nuclei and collagen Solutions required: A. Haematoxylin 5% in absolute alcohol . . . . . . . . 20 ml. 218 SECTION TWO Ferric chloride (hydrated) io% aqueous . . . . . . 8 ml. Iodine solution (i gm. iodine, 2 gm. KI, 50 ml. water) . . 8 ml. Note: Solution A deteriorates after twenty-four hours. B. Ferric chloride hydrated 2% aqueous. C. Van Gieson stain. Tissues should be fixed in Zenker or in 10% formalin: if the former is used mercurial precipitates are removed by the iodine in the staining solution and it is not, therefore, necessary to treat the sections or tissues with iodine before staining. Technique: Paraffin wax, Celloidin or L.V.N, may be used for embedding. 1. Sections are brought down to distilled water; then im- mersed in Solution A for one quarter to one hour until quite black. 2. Differentiate for a few minutes in Solution B, controlling by examination in water under the low-power objective. 3. Wash with tap water; then immerse in 95% alcohol to remove iodine. 4. Wash in tap water for five minutes; then counterstain in Van Gieson for three to five minutes. 5. Differentiate in 95% alcohol; then dehydrate. 6. Paraffin sections are cleared in xylol ; Celloidin or L.V.N, in terpineol (after 95% alcohol). 7. Mount in Cristalite or in balsam. Results: Elastic fibres, intense blue-black to black. Nuclei, blue to black. Collagen, red. Other tissue elements, yellow. 219 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES WATER BLUE - ORCEIN - SAFRANIN For demonstrating epithelial fibres Solutions required: A. Water blue . . . . . . i gm. Orcein . . . . . . . . 0-75 gm. Glycerine . . . . . . . . 20 ml. Absolute alcohol . . . . 50 ml. Acetic acid 5% . . . . . . 100 ml. B. Eosin 1-25% alcoholic. C. Hydroquinone 1% aqueous. D. Safranin O, aqueous 1%. E. Potassium dichromate 0-5%. Technique: 1. Specimens of skin are fixed in 10% formalin and embedded either in paraffin wax or in Celloidin. 2. Bring sections down to distilled water; then stain for ten minutes in a mixture consisting of: Solution A . . . . . . 10 ml. Solution B . . . . • • 3 ^^ Solution C . . . . • • 3 rnl. 3. Wash well in distilled water. 4. Stain for ten minutes in the safranin solution. 5. Wash thoroughly in distilled water. | 6. Immerse in 0-5% potassium dichromate solution from ten to thirty minutes. 7. Wash in distilled water; dehydrate in absolute alcohol ; then clear in oil of bergamot. 8. Examine under the microscope; then if necessary differ- entiate alternatively with absolute alcohol and oil of bergamot until the depth of the safranin stain has been reduced. 9. Mount in balsam or in Cristalite. Results: Epithelial fibres are stained red, while the nuclei are pale violet ; 220 SECTION TWO plasmasomes, red; cytoplasm, blue to violet; granules of the neutrophil leucocytes, sky blue; elastic fibres, red; collagen fibres, blue. WEIGERT - FRENCH ELASTIN STAIN (Moore's modification) This modification, which is due to G. W. Moore, of the Central Histological Laboratory, Archway Hospital, London, gives greater selectivity than either Sheridan or Weigert elastin stains, and consistently excellent results are obtained provided the stain is properly prepared. Solutions of Moore's elastin stain will keep for several years without deterioration. The dry stain requires time and great care for its preparation; the majority of workers will, no doubt, wish to purchase the stain ready for use, but for those who have the time and prefer to pre- pare the stain themselves, the method is given below. Preparation of the dry stain: A. Ferric chloride, anhydrous, A.R. grade . . . . . . . . 30 gm. Distilled water . . . . . • 65 ml. Dissolve; then make up the vol- ume to 100 ml. with distilled water. Note: This solution must be freshly prepared. . Crystal violet » • • • • 2-5 gm. Basic fuchsin * m • • • 2-5 gm. Dextrin • • « • i-ogm. Resorcin, pure 1 • • • • 4 lo-o gm Distilled water 1 • • • • 500 ml. Note: The water must be heated to about 95° C. in a large evaporating basin, the dyes and dextrin added and stirred until dissolved. The resorcin is then added and the solution raised to boiling point with constant stirring. 221 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. When Solution B begins to boil, add 62 ml. of Solution A slowly, in small portions at a time over a period of five minutes, with constant stirring. 2. Continue with the boiling and stirring for a further period of two or three minutes until a coarse precipitate is obtained. 3. After cooling filter by means of a Buchner funnel and flask attached to a suction pump. 4. Wash the precipitate with distilled water until the runnings are colourless and the filtrate a clear azure blue; this usually requires 8 to 10 litres distilled water. 5. The preparation is then dried overnight in an incubator, after removing the filter paper. Preparation of the staining solution : 1. The dried elastin stain is now dissolved in 550 ml. absolute alcohol plus I ml. HCl in a i -litre flask, the neck of which is plugged lightly with cotton-wool. Solution is effected by boiling gently for about thirty minutes on a water bath or electric hot- plate. 2. Cool; filter; add 19 ml. concentrated HCl; then shake well and allow to stand for at least twenty-four hours. Staining technique: 1. Sections are brought down to distilled water; then treated with 0'5% aqueous potass, permanganate for five minutes. 2. Rinse and bleach with 5% aqueous oxalic acid; then wash in running water. 3. Transfer to elastin stain for at least two hours at 37° C. or for one half to one hour in an oven at 60° C. 4. Blot and treat with absolute alcohol for three to five minutes. 5. Rinse with water and counterstain for three minutes with neutral red (Jensen). 6. Rinse and differentiate the neutral red for a few seconds in absolute alcohol. 7. Rinse in distilled water, then pour on 0-5% picric acid aqueous and wash off immediately with running water. 222 SECTION TWO 8. Blot; dehydrate in absolute alcohol; clear in xylol and mount. Results: Elastic fibres, blue-black. Nuclei, red. Erythrocytes and muscle, yellow. Notes: Best results are obtained after the stain has been kept in stock for several weeks, when it becomes perfectly selective and remains so indefinitely. The use of Coplin's jars, which can be " topped-up ** occa- sionally to make good loss by evaporation, is to be recommended. The picric acid gives a beautiful contrast to the neutral red and enhances the appearance of the elastic fibres by causing them to stand out against a neutral background. Care must be taken when using it, however, as overstaining tends to give the red nuclei an unpleasant brownish tinge. It is perhaps advisable, until the technique has been mastered, to use neutral red only, ensuring that it is properly differentiated in absolute alcohol. WEIGERT - PAL TECHNIQUE For myelin sheaths in brain and spinal cord and for peri- pheral nerves and ganglia Solutions required: A. Weigerfs rapid fixative : Potass, dichromate 5gm. Fluorchrome powder 2gm. Distilled water 100 ml. Dissolve by heat; cool and filter • B. Haematoxylin io% in absolute alcohol . . 10 ml. Absolute alcohol 90 ml. C. Saturated lithium carbonate, aqueous 7 ml. Distilled water 93 aaaI. D. Potass, permanganate 0-25% E. Oxalic acid i% . . 50 ml. Potass, sulphite, anhydrous i%. . 50 ml. 223 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Slices of the material 2 to 5 mm. thick are fixed in 10% formalin ; then transferred to Solution A for four to seven days. 2. Wash in running water for several hours; then dehydrate and embed in Celloidin, L.V.N, or in paraffin wax, or cut frozen sections (in which case 2% ammon. bromide should be added to the formalin fixing solution). 3. Sections 10 to 20 fi in thickness are stained twenty-four to forty-eight hours in a freshly prepared mixture consisting of i vol- ume Solution B and 9 volumes Solution C. 4. Immerse for one half to three minutes in Solution D ; then rinse in distilled water. 5. Differentiate in Solution E for one half to three minutes or until the white matter is blue-black, and the grey matter almost colourless. 6. Counterstain with safranin 1% aqueous, if desired, for one half to two hours according to thickness of sections. 7. Wash thoroughly in water. 8. Dehydrate ; clear and mount. Results: Myelin sheaths, blue-black. Myelinated fibres, black or blue- black. Grey matter, white or slightly yellow. Other structures, unstained (unless a counterstain has been used). WOOL GREEN - HAEMATOXYLIN - PONCEAU S For connective tissue and muscle Solutions required: A. Picric acid, saturated in 70% alcohol. B. Weigert's haematoxylin A. C. Ponceau S 1% in 1% acetic acid aqueous. D. Weigert's haematoxylin B. E. Wool Green S 1% in 1% aqueous acetic acid. F. Acetone and xylol, equal volumes of each. 224 SECTION TWO Technique: 1. Immerse sections in the picric acid solution for two minutes. 2. Wash thoroughly in running tap water. 3. Stain for five to seven minutes in Weigert's haematoxylin A. 4. Rinse in water and stain for three to five minutes in the Ponceau S solution. 5. Wash in water. 6. Immerse in Weigert's haematoxylin B. 7. Wash thoroughly in water. 8. Stain for three to five minutes in the wool green solution. 9. Decolorize for two minutes with 1% acetic acid. ID. Pour off excess acid; rinse well in distilled water; drain and blot carefully. 1 1 . Rinse well with acetone. 12. Rinse with two washings of Solution F. 13. Clear in xylol and mount. Results: Muscle and cytoplasm, red. Connective tissue and basement membranes, green to dark blue. WRIGHT'S STAIN For general differentiation of blood corpuscles; for malarial parasites; trypanosomes, etc. This stain is extensively used in America instead of Leishman stain which is preferred by British workers. Solutions required: Formol-saline^ neutral^ buffered: A. Formalin (40% formaldehyde) . . 100 ml. Sodium chloride, A.R. . . . . 8-5 gm. Distilled water . . . . . . i litre Acid sodium phosphate, mono- hydrate . . . . . . • . 4 gm. Anhydrous disodium phosphate 6-5 gm. 225 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES B. Wright's stain. C. Acetic acid, o-o8% aqueous. Technique: 1. Fix pieces of tissue in Solution A for sixteen to forty-eight hours. 2. Dehydrate in the usual ascending grades of alcohol; clear and embed in paraffin wax. 3. Fix sections, not exceeding 5// in thickness to slides; remove wax with xylol ; pass through descending grades of alcohol down to neutral distilled water. 4. Stain for three to five minutes in a freshly prepared mixture consisting of one volume of Wright's stain and two volumes of neutral distilled water, in a stoppered staining jar. 5. Rinse with neutral distilled water. 6. Differentiate with the acetic acid solution, controlling by examination under the microscope, until the protoplasm of the cells is pink, and only nuclei are blue. 7. Wash with neutral distilled water. 8. Dehydrate quickly with absolute alcohol; clear in xylol; mount in Cristalite. Results: Erythrocytes, yellowish red. Polymorphonuclears, dark purple nuclei, reddish violet granules, pale pink cytoplasm. Eosinophiles, blue nuclei, red to orange-red granules, blue cytoplasm. Baso- philes, purple to dark blue nuclei, dark purple to black granules. Lymphocytes, dark purple nuclei, sky blue cytoplasm. Platelets, violet to purple granules. Malarial parasites and Leishmanial chromatin, red; cytoplasm, blue. Trypanosomes : chromatin, red. Note: The timing of the staining either before or after dilution may be altered to suit individual requirements. Staining effects similar to Giemsa are obtained by staining for ten minutes in Wright's stain diluted with four times its volume of distilled water buffered to pH 6-5. 226 SECTION 3— BOTANICAL METHODS (Normal and Infected Tissues) (a) GENERAL TREATMENT OF TISSUES Material must be killed and fixed immediately it is collected to ensure that tissues are preserved with as life-like an appearance as possible. Small organisms such as unicellular algae, filamentous fungi, etc., may be placed directly into the killing fluid. Large objects must be cut at once into pieces not exceeding 5 mm. in any one direction, and care must be taken to avoid rough handling and pressure. There are a great number of killing and fixing fluids to choose from and the reader should refer to the chapter on fixatives in this book as well as to standard text-books on botany such as Johansen or Chamberlain. The best general killing fluid, however, is 70% alcohol containing o-i% glacial acetic acid; material should be immersed in this fluid from ten minutes to an hour. After killing and fixing, material may be preserved, if it is not required for immediate examination, in 70% alcohol containing 20% glycerine, or in 10% formalin buffered to pH 7-0. Delicate material should be stored in 85% alcohol containing 20 ml. glycerine per 100 ml. while tough hard material is best stored in 50% alcohol containing 20 ml. glycerine per 100 ml., but here again the reader is referred to standard text-books on botanical technique. Material taken from the fixing and killing fluid or from the preserving fluid should be washed well in running water. Tissues may be dehydrated, cleared and embedded, as in the case of animal tissues. Alternatively, sections may be cut, without dehydration, clearing and embedding, either by means of a hand microtome, or by placing thin slices of the material in a slit cut in a piece of elder pith and cutting sections " freehand '* with the razor edge facing away from the operator. In cutting freehand sections, the razor should be kept wet with 10% aqueous glycerine. Glycerine (pure or diluted with water), or glycerine jelly should be employed for mounting temporary preparations after staining, while the preparation of permanent slides may be carried out in 229 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES the same manner as for sections of animal tissues, except that dehydration should be more gradual in order to avoid violent diflFusion and consequent shrinkage and distortion of delicate tissues; the alcohols used should be graded 50%, 70%, 80%, 90%, 95% and absolute. (b) MISCELLANEOUS MICROCHEMICAL TESTS Note: In the following tests fresh material, in general, should be employed and thick sections, up to about 40 or 50/^ are prefer- able to thin ones, particularly if the substances under test occur in small proportions. Aldehydes Solution required: Diphenylamine 1% in concen- trated sulphuric acid. Technique: 1. Sections of strictly fresh tissue are placed directly into a drop of the reagent on a slide. 2. Heat gently until a green coloration appears. 3. Continue the heating for about five minutes. Results: If the green colour persists, formaldehyde is indicated but if the green turns to red, aldehydes, other than formaldehyde, are present in the tissue. Aleurone This occurs in the seeds of Poaceae Solution required: Eosin 2% in saturated alcoholic picric acid. Technique: I. Sections are immersed in a few drops of the reagent on a slide and examined under the microscope. 230 SECTION THREE 2. As soon as the ground substance of the aleurone grains appear blue, add a few drops of absolute alcohol, when the globules should be colourless and the crystals a yellowish green. 3. Remove excess alcohol; clear in clove oil and mount in balsam. Results: Protein (aleurone) crystals, yellow. Globules, pink. Ground substance, dark red. Amygdalin This occurs in the seeds of Amygdalus; Pyrus Crataegus and related genera, as well as the leaves of Prunus laurocerasus Solutions required: A. Picric acid, saturated, aqueous. B. Sodium carbonate 10% aqueous. C. Potassium hydroxide 4% in 70% alcohol. D. Ferrous sulphate 2-5% aqueous (prepared without heating). E. Ferric chloride 20% aqueous. F. Hydrochloric acid, cone. . . i volume Distilled water . . . . • • 4 volumes Technique (a): 1 . Immerse sections in the picric acid solution for half an hour, on slides. 2. Wash with water ; then add one drop of the sodium carbon- ate to a section. Result: A red coloration indicates the presence of amygdalin, and this is confirmed by the odour of hydrocyanic acid (almond odour), which is a decomposition product upon which tests for amygdalin are based. R 231 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique (b): 1. Place sections in the sodium hydroxide (Solution C) for three to five minutes. 2. Mix equal volumes of the ferric sulphate and ferric chloride (Solutions D and E above) in a test-tube or small beaker and heat to boiling. 3. Immerse sections in the above mixture and leave them therein for five to ten minutes. 4. Place sections on a slide in a drop of the diluted hydrochloric acid (Solution F above). Results: A deep blue precipitate (Berlin blue reaction) indicates the presence of hydrocyanic acid which is indicative of amygdalin. Amylodextrin This product, which is intermediate between maltose and starch, is present in solution in storage organs where starch is hydro- lysed. Its presence is indicated by the red coloration it produces with Gram's iodine solution Anthocyanin This occurs in the petals of the vast majority of blue and red flowers; such flowers owe their colour to anthocyanin Solutions required: A. Glacial acetic acid. B. Ammonia solution. Technique: • 1. Place petals in a little neutral distilled water on slides. 2. Cover with thick coverslip (No. 2 or No. 3) and press out the coloured liquid. 3. Run in a drop of glacial acetic acid under the coverslip. 232 SECTION THREE 4. Take another slide and repeat the process but run in a drop of strong ammonia solution instead of acetic acid. Results: Anthocyanin is red in acid solution and blue violet to green in alkaline solution. Arbutin This occurs in Ericacae and Pyrolacae Solutions required: A. Nitric acid 10% B. Ammonia solution. C. Ferric chloride 5% aqueous. Technique {a): 1. Place sections in the nitric acid solution. 2. Cover with a coverslip and examine under the microscope i mmediately. Results: Cells containing arbutin assume a dark orange colour, which rapidly changes to yellow which slowly disappears altogether. Technique (b): This depends upon the principle that arbutin is converted, on hydrolysis, to glucose and hydroquinone. 1 . The dry sections are placed in a drop of water on a slide and heated gently for two or three minutes, when the arbutin sublimes in crystals. 2. Add a drop of ammonia solution when the arbutin crystals assume a rich brown colour. 3. Repeat the process adding a drop of ferric chloride solution instead of ammonia : a pale green colour confirms the presence of arbutin. 233 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Asparagine This occurs widely in the plant kingdom, but is most readily demonstrated in etiolated seedlings of Lupins and tubers of Dahlia Solution required: Cupric acetate 5% aqueous. Technique: 1. Place strictly fresh sections in a drop of the cupric acetate on a slide and leave therein for ten to twenty minutes. 2. Add absolute alcohol slowly, a drop at a time, whilst examin- ing under the microscope, until ultramarine spaerocrystals of copper asparagine become visible, indicating the presence of asparagine in the tissue. Calcium Solution required: Oxalic acid 2% aqueous. Technique: 1. Sections are placed directly onto a slide and flooded with the oxalic acid solution and left for half an hour exposed to the air. 2. Pour off some of the liquid ; then add a coverslip. 3. Pipette a drop of absolute alcohol along one edge of the coverslip, so that the alcohol is drawn under the coverslip by capillary attraction. Examine under the microscope. Results: Calcium, if present, will be indicated by the small but easily visible crystals of calcium oxalate. Calcium Oxalate Solutions required: A. Cupric acetate, saturated, aqueous. B. Ferric sulphate . . . . • • 5 gn^« Acetic acid 20% aqueous . . 100 ml. 234 SECTION THREE Technique: 1. Sections are placed directly into a drop of the cupric acetate solution on a slide and left therein for about ten minutes. 2. Examine under the microscope and if calcium oxalate crystals are present they will have dissolved and the oxalic acid diffused into the intracellular spaces where cupric oxalate is formed. 3. To test for the dissolved oxalate, add a few drops of the ferric sulphate (solution B) and examine under the microscope. The appearance of yellow ferrous sulphate crystals confirms the pres- ence of calcium oxalate, in the tissue. Callose Solution required: Lacmoid I % alcoholic .. .. o-i ml. Distilled water . . . . 25 ml. Technique: 1. Immerse sections in the lacmoid solution for about fifteen minutes, when callose, if present, is stained a brilliant blue. 2. Mount in a drop of glycerine on a slide and examine under the microscope. The following solubility tests may be employed to distinguish callose from other membrane substances: {a) Soluble in copper oxide ammonia, while cellulose and hemi- celluloses are insoluble. {b) Swells but is insoluble in solutions of ammonia, sodium carbonate, potassium carbonate, whilst pectic acid is soluble in these reagents. {c) Soluble in glycerine at 280° C. whilst cellulose and chitin are insoluble. Carotin Solution required: Potassium hydroxide 20% in absolute alcohol. Technique: I . Place sections of fresh young green leaves in the potassium 235 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES hydroxide solution in a stoppered jar and leave in the dark until the chlorophyll is extracted. 2. Remove the sections from the jar and wash them for ten hours in running water. 3. Transfer to a large volume of distilled water and leave therein for an hour, afterwards transferring to a fresh lot of distilled water for a further period of one hour. 4. Transfer to slides ; mount in glycerine. Results: Carotin, if present, appears as orange-red crystals. Xanthophyll will appear after two or three days as yellow crystals. Cellulose Solutions required: A. Gram's iodine. B. Sulphuric acid 75%. Technique: 1. Place sections in a drop of Gram's iodine solution on a slide. 2. Cover with a coverslip; examine under the microscope, taking careful note of the location of the blue coloration. 3. Place a drop of the sulphuric acid solution along the edge of the coverslip and observe the swelling of the cellulose membranes as the sulphuric acid seeps under the coverslip, hydrolysing the cellulose to a colloid substance known as hydrocellulose. Note: As certain other plant substances give a blue colour reaction with iodine it is important to note the locality of any blue colour which appears prior to the hydrolysation with the sul- phuric acid. Chitin This is said to occur principally in the higher fungi. Solution required: A. Potassium hydroxide . . . . 78 gm. Distilled water . . . . . . 68 ml. 236 SECTION THREE Technique: 1 . Heat the potassium hydroxide solution in an open beaker to boiling point. 2. Place the sections in the beaker which should now be covered with a clock glass, and continue the boiling for half an hour. 3. Remove the sections and wash in 90% alcohol. 4. Treat the section with Gram's iodine solution. Results: Chitin, if present in the tissue, will be indicated by a reddish- violet colour. Chlorides Occur in roots of Dauctis carota and Beta; Primula ohconica; solanum Solutions required: A. Silver nitrate 5% aqueous. B. Nitric acid 1-5% aqueous. Technique: 1 . Sections, which must be cut with a scrupulously clean razor, are placed in a drop of the silver nitrate solution on a slide. 2. Examine, without a coverslip, under the microscope, when the silver chloride precipitate will appear black. Note: The presence of chlorides in the tissue is indicated by a precipitate which is white to the naked eye. 3. By means of glass needles transfer the sections to a drop of the diluted nitric acid solution on another sHde. 4. Examine under the microscope when it will be observed that the acid clears the sections sufficiently to allow localization of the reaction. 5. Repeat stage 3 (above); add a few drops of ammonia solu- tion to the slide until the precipitate just dissolves. 237 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 6. Examine again after an hour when the precipitate will re- appear in crystalline form. Chlorophyll Solutions required: A. Potassium hydroxide 20% in pure methyl alcohol. Technique: 1 . Place sections directly onto a slide and add two or three drops of ether. 2. Add a few drops of the potassium hydroxide solution. Results: Chlorophyll immediately turns brown, afterwards changing back to green again. Formic Acid Solutions required: A. Mercuric chloride 1% aqueous. B. Hydrochloric acid, cone. C. Potassium hydroxide 1% aqueous. Technique: 1. Place fresh sections on slides; flood with the mercuric chloride solution and heat on a water bath for an hour. 2. Pour off excess mercuric chloride solution and wash with dis- tilled water which has been acidified by the addition of i ml. hydro- chloric acid cone, per 100 ml. 3. Place sections on slides; add one drop of 1% potassium hydroxide, and examine. Results: Where formic acid is present, the cells are blackened. 238 SECTION THREE Glutathione Solutions required: A. Acetic acid i%. B. Ammonium sulphate, saturated, aqueous. C. Sodium nitroprusside 5% aqueous. Technique: 1. Place thin sections of strictly fresh tissue on slides. 2. Flood with 1% acetic acid and heat gently till vapour rises. This is most conveniently done by placing the slides over a corner of a tripod and applying the heat by means of a very small bunsen flame which should be held some distance away from the underside of the slide. 3. Transfer to a watch glass and rinse in the ammonium sul- phate solution. 4. Immerse in a mixture consisting of 0-5 ml. of the sodium nitroprusside solution and 5 ml. of saturated ammonium sulphate, in a watch glass. 5. Agitate gently but thoroughly by rocking the watch glass for a few minutes. 6. Whilst watching the sections closely, add i ml. of ammonia solution. Results: If glutathione is present the cells will assume a red colour, which usually lasts only a second or so, with the addition of the ammonia solution. Inulin Occurs in bulbs of Dahlia variabilis^ etc. Solutions required: A. Thymol 15% in absolute alcohol. B. Chloral hydrate . . . . . . 10 gm. Distilled water . . . . • • 4 ml- 239 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Fresh tissues are fixed in 70% alcohol for three or four days. 2. Place sections in alcohol on slides and add a drop of the chloral hydrate solution. 3. Examine under the microscope and observe concentric layers of inulin crystals, if present. 4. Add one drop of the thymol solution followed by concen- trated sulphuric acid. Results: On addition of the last two reagents inulin crystals immediately become red, dissolving after a minute or so. Iodine Solutions required: A. Starch 1% aqueous suspension. B. Potassium nitrite 20% aqueous. C. Hydrochloric acid cone. . . i ml. Distilled water . . . . . . 19 ml. Technique: I . Place fresh sections in a watch glass containing 2 or 3 ml. of the starch suspension together with three or four drops each of Solutions B and C. Results: Iodine is indicated if the starch is coloured blue. Iron Note: The use of iron or steel instruments in the following technique should be avoided. The section razor, which must, however, be used, should be scrupulously clean. Solutions required: A. Hydrochloric acid concentrated . . 2 ml. Distilled water . . . . . . 98 ml. Potassium ferrocyanide 10% . . 2 ml. B. Alum carmine. 240 SECTION THREE Technique: 1 . Sections of fresh material are taken from distilled water and immersed in Solution A (above) for one half to one hour. 2. Wash well with several changes of distilled water. 3. Stain the nuclei with alum carmine solution for a few min- utes , then wash well with distilled water. 4. Dehydrate through ascending grades of alcohol as usual, clear in xylol ; mount in D.P.X. or Cristalite. Results: Nuclei, red. Iron (if present), blue. Lecithin Solutions required: A. Scarlet R (Botanical). B. Delafield haematoxylin. Technique: 1. Fat is removed from sections by immersion overnight in acetone, in a stoppered jar or well-corked tube. 2. Fix sections to slides; rinse in two changes of pure acetone. 3. Stain the scarlet R solution for about ten to fifteen minutes. 4. Wash quickly with 70% alcohol. 5. Wash in distilled water. 6. Counterstain with Delafield haematoxylin for two to ten minutes. 7. Blue and wash in tap water. 8. Mount in glycerine or Aquamount. Results: Lecithin (if present), red. Nuclei, blue. 241 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Nitrates Solution required: Diphenylamine . . . . . . i gm. Sulphuric acid, cone. . . • • 75 "il- Distilled water . . . . • • 25 ml. Add the sulphuric acid cautiously to the water in small portions (about 5 ml.) at intervals in a 250-ml. conical flask, swirling the contents round gently to ensure thorough mixing. Do not on any account add the water to the acid as this will result in the acid flying back and causing serious injury to the face and hands , clothing , etc. After the diluted acid has cooled somewhat, add the diphenylamine. Technique: 1. Place sections on slides under coverslips. 2. Place two drops of the diphenylamine solution along one edge of the coverslip. Results: If nitrates are present a deep blue colour develops as the diphenylamine-sulphuric acid solution seeps under the coverslip and comes into contact with the section. After a few minutes, the section disintegrates and the colour changes to light brown. Pectic Substances Solution required: A. Ruthenium red, 0-02% aqueous Technique: 1. Stain with the ruthenium red for thirty minutes. 2. Mount in glycerine on a slide. Results: Pectic substances are stained red. 242 SECTION THREE Phosphates Solution required: Ammonium molybdate io% in concentrated nitric acid. Technique: Place sections on a slide in a drop of the reagent, and examine under the microscope. Results: Phosphates are indicated by small black-bordered yellow drops, which develop into spaerocrystals and afterwards into cubes and octahedrons. Phjrtosterol Solution required: Gram's iodine. Technique: 1. Place thick sections on slides and cover with concentrated sulphuric acid. 2. If phytosterol is present sections will assume a red color- ation. Add a drop of Gram's iodine solution and mix by rocking the slide backwards and forwards gently. Results: The presence of phytosterol is confirmed when the colour changes from red, after the addition of the iodine, to violet ; then blue and finally to yellowish red or brown. Potassinm Solution required: Sodium Cobalt Nitrite. Sodium nitrite . . . . • • 7 gn^« Cobalt nitrate . . . . • • 4 g"^- Distilled water . . . . . . 13 ml. Glacial acetic acid . . . . 2 ml. 243 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: I. Sections of fresh tissue are placed in a drop of the above reagent on a sHde and examined under the microscope. Results: The appearance of fine yellow crystals of potassium cobalt nitrite indicates the presence of potassium. Proteins Solutions required: A. Potassium ferrocyanide . . . . o-8 gm. Acetic acid . . . . . . loo ml. B. Ferric chloride aqueous 5%. Technique: 1. Immerse sections of fresh material in Solution A for an hour. 2. Rinse quickly with 60% alcohol. 3. Add a few drops of 5% ferric chloride. Results: A blue coloration indicates proteins. Saponin Method (a) Place a drop of concentrated sulphuric acid on a section of fresh material. If saponin is present in the tissue the section immediately assumes a yellow colour which changes to red after about half an hour, and later to violet or bluish green. Method (b): Solutions required: A. Barium hydroxide, saturated aqueous. B. Calcium chloride 5%. C. Potassium dichromate 10% aqueous. 244 I SECTION THREE Technique: 1. To locate the sites of saponin immerse sections in the barium hydroxide solution for sixteen to twenty-four hours ; then examine under the microscope and observe the insoluble colourless com- pound formed by the interaction of saponin and barium. 2. Wash well with the calcium chloride solution. 3. Cover with the potassium dichromate solution and watch the reaction under the microscope. Results: The insoluble substance first formed between the barium and saponin is broken down, the barium uniting with the chromium, forming barium chromate, which is identified by its yellow colour. Cells containing tannin assume a rich brown colour during the reaction. Sodium Solution required: Uranium acetate, saturated aqueous. Technique: 1. Sections are placed in a few drops of the reagent on a slide. 2. Add one drop of hydrochloric acid to the preparation. 3. Place uncovered slides in a desiccator to facilitate the slow evaporation of the reagents. 4. Examine at hourly intervals over a maximum period of eight hours. Results: Pale yellow rhomboidal or tetrahedral crystals (sodium uranium acetate) indicate the presence of sodium in the tissue. Note: The presence of magnesium is indicated by large rhom- boidal crystals, with this technique. 245 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Sulphates Solutions required: A. Benzidine hydrochloride. . Hydrochloric acid, cone. Distilled water B. Hydrochloric acid, cone. Distilled water I gm. 3 ml. 97 ml. 10 ml. 90 ml. C. Barium chloride 10%. Technique {a): Sections of fresh tissue are placed in a few drops of the benzi- dine solution on a slide and examined under the microscope. Results: Scales or small glistening needles (benzidine sulphate) indicate the presence of sulphates. Technique (b): For cereal seeds and other tissues which contain fat. 1. Immerse the material overnight in acetone in a stoppered jar or well-corked tube, to remove the fat. 2. Transfer to slides; wash in two changes of pure acetone; then allow the sections to dry on the slides. 3. Add one drop each of Solutions B and C (above) to the slide and examine under the microscope, under a high-power objective. Results: A granular precipitate (barium sulphate) indicates the presence \ of sulphates. Tyrosine Solutions required: Sodium molybdate 1% in sul- phuric acid, cone. Technique: 1. Place sections on slide and cover with absolute alcohol. 2. Allow the alcohol to evaporate completely. 246 SECTION THREE 3. Cover the section with a few drops of the sodium molybdate and warm gently for a few minutes. Results: A deep blue colour turning to violet after a few minutes indicates the presence of tyrosine. (c) STAINING TECHNIQUES ACID FUCHSIN - AURANTIA For di£Ferentiating between bacteria and mitochondria in sections of infected tissue Solutions required: A. Chromic acid 1% aqueous . 50 ml. Potass, dichromate 1% aqueous. . 50 ml. Neutral formalin . . . 8 ml. B. Acid fuchsin . 2gm. Aniline water ID ml. C. Aurantia . . . 0-5 gm. Alcohol 70% . 100 ml. D. Phosphomolybdic acid . . . I gm. Sodium hydroxide 1% aqueous. . 10 ml. Distilled water 100 ml. E. Polychrome methylene blue (Un na). Technique: 1 . Kill and fix for twenty-four hours in Solution A. 2. Wash for twenty-four hours in running water; then de- hydrate and embed. Sections should be cut as thinly as possible. 3. After removing the paraffin wax from slides they are dipped into a very thin solution of Celloidin in equal parts of absolute alcohol and ether; then passed through absolute, 90%, 70% alcohol to distilled water. 4. Stain in Solution B, heated to about 80° C, for a few minutes, then wash in running water. 247 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 5. De-Stain for a few seconds in Solution C ; then wash in water. 6. Immerse in Solution D for a few minutes; then rinse in water. 7. Stain for a few minutes in Solution E, then wash with water. 8. Dehydrate very quickly with 95% and absolute alcohol; clear in xylol and mount. Results: Bacteria, deep violet blue. Mitochondria and plastids, red. ACID RUBIN - AURANTIA - TOLUIDINE BLUE (KuU's Stain) For starch grains and mitochondria in plant tissues Solutions required: A. Acid rubin 1% aqueous. B. Aurantia 5% in 80% alcohol C. Tannic acid 2% aqueous. D. Toluidine blue 1% aqueous. Technique: 1. Fix material in Regaud's fluid. 2. Stain for five minutes in solution A heated to about 60 or 70° C. 3. Differentiate with solution B, controlling by examination under the microscope. 4. Wash in water. 5. Immerse in solution C for 20 minutes. 6. Wash well in water. 7. Stain for five minutes in solution D. 8. Pour off excess stain and rinse with 70% alcohol. 9. Differentiate in 90% alcohol. 248 SECTION THREE 10. Dehydrate with absolute alcohol. 1 1 . Clear in xylol, and mount. Results: Mitochondria are stained red, while starch grains are blue. Note: This is an adaptation of one of Volkonsky's techniques. Reference: Milovidov (1928), Arch. Anat. Micros., 24, 9. ANILINE HYDROCHLORIDE A simple and rapid method of demonstrating lignified tissues Solution required: Aniline hydrochloride 10% aqueous, freshly filtered before use. Technique: 1. Place sections on slides and cover with a few drops of the aniline hydrochloride solution, and allow this reagent to act for about five minutes. 2. Pour off excess ; place a drop of glycerine on the section and cover with a coverglass. Results: Lignified tissues are stained yellow while the other tissues remain unstained. BASIC FUCHSIN, AMMONIACAL For lignified walls and cutin Solution required: A. Basic fuchsin, 10% alcoholic. B. Strong ammonia solution . . 25 ml. Note: The fuchsin solution is added drop by drop to the ammonia until a yellowish colour is produced. 249 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES The fuchsin is decolorized by the ammonia and if too much fuchsin is added, then a few more drops of ammonia must be added so that the final product is pale yellowish in colour. Technique: 1. Immerse sections in the solution in a stoppered jar for five to ten minutes. 2. Transfer sections to watch glasses containing absolute alcohol, for about five minutes, until the alcohol takes on a pink coloration. 3. Transfer sections to a fresh lot of alcohol and leave therein for about five to ten minutes. 4. Rinse and dehydrate in a fresh lot of absolute alcohol. 5. Clear in clove oil. 6. Mount in Canada balsam. Results: Lignified walls and cutin, intense red; remainder, colourless. BORAX CARMINE (Grenacher) For bulk staining prior to sectioning, and for small whole mounts Solutions required: A. Borax carmine, alcoholic (Grenacher). B. Alcohol 70% . , . . . . 100 ml. Hydrochloric acid, concentrated 0-25 ml. Technique: 1. Immerse material in the carmine solution from one hour to four days, according to the bulk and nature of the material, until sufficiently stained. 2. Rinse with and immerse in acid alcohol (Solution B, above) until clear. 3. Wash with 70% alcohol. 4. Dehydrate with 95% alcohol, followed by absolute. 250 SECTION THREE 5. Clear in xylol. 6. Embed in paraffin wax. 7. Cut sections and mount on slides. 8. De-wax with xylol. 9. Mount in Canada balsam in xylol. Note: For whole mounts, after clearing in xylol (Stage 5), mount in balsam. Results: Nuclei are stained deep red, while cytoplasm is pink. CHLORAZOL AZURINE A simple double stain, non-fading, and particularly suitable for elementary class work Solutions required: A. Formaldehyde 40%. . .. .. 5 ml. Acetic acid 50% . . . . . . 14 ml. Absolute alcohol . . . . • . 63 ml. Distilled water . . . . . . 20 ml. B. Magnesium sulphate, crystals, 6% aqueous. C. Chlorazol azurine, saturated aqueous. D. Equal volumes of Solutions B and C. Heat the mixture to 80° C. and stir well for five or ten minutes, taking care that this temperature is not exceeded by more than a few degrees. Technique: 1 . Sections of the material, fresh from the field, are transferred to solution A. Alternatively material may be stored in this fixative for several days before sectioning. 2. Take sections down to water. 3. Immerse sections in solution D (the solution should be shaken well immediately before use) in a jar or tube overnight, or for at least eight hours. 251 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 4. Wash well with tap water. 5. Wash quickly in 70%, 90% and 95% or absolute alcohol. 6. Mount directly in Michrome mountant or in Euparal. Result: Non-lignified cell walls, blue, Lignified cells, violet to red. Bark cells, orange. Herbarium Specimens Solution required: Solution D, as above. Technique: 1. Soak or boil the specimens in water. 2. Cut sections and stain directly with solution D, which has been well shaken immediately before use. Note: Prolonged treatment in the fixatives employed bleaches the specimens sufficiently for the purpose of staining with chlorazol azurine ; otherwise bleaching is not recommended. Reference: Armitage, F. D., ^. Roy. Micr. Soc.y 535, 826, i. / am indebted to Mr. F. D. Armitage^ F.R.M.S.y of The Labora- tory, Green End Road, Boxmoor, Herts, England, for information he has given me regarding his use of this stain. CHLORAZOL BLACK A non-fading, general-purpose stain, which may be used for whole mounts as well as for sections. The stain requires no mordanting nor differentiation Nuclei and chromosomes are stained black, cytoplasm and secreted products grey, by this stain, which has also been found useful for infected plant tissues. Solution required: Chlorazol black, saturated in 70% alcohol. 252 i SECTION THREE Technique: 1. Fix tissues in Bouin or Flemming and embed in paraffin wax. 2. Stain in a freshly prepared, unfiltered, alcoholic solution of chlorazol black (as above) for five to ten minutes. 3. Drain off excess stain; dehydrate; clear in xylol and mount. Results: Vascular plant: Cell wall, jet black; cytoplasm, greyish green; nuclei, yellowish green; nucleoli, deep amber to dark green. Fern leaf: Cell wall, intense black; epidermis walls, heavy black; cytoplasm, light amber; nuclei, green; nucleoli, dark green; plastids, grey; suberized walls of midrib, dark amber; veins, dark amber. Notes: (a) The stain may be incorporated with Lactophenol. {h) Benzyl alcohol may be used as a solvent of the stain, in which case the results are somewhat different. {c) If it is desired to differentiate the stain dilute " Milton '* (a proprietary antiseptic) may be used. CHLORAZOL PAPER BROWN, B For plant tissues Material may be stained overnight, but in many cases, e.g. where the delicate cell contents are not germane to investigation, the equivalent depth of staining will be produced by boiling for one to two minutes in the staining solution. By employing the boiling technique finished slides can be obtained in five minutes. Solution required: Chlorazol Paper Brown,^ B, satur- ated aqueous. Technique: I. Sections are stained overnight in a saturated aqueous solution (about 3%) of the stain, or by boiling in this staining solution for one to two minutes. 253 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 2. Differentiate in i% nitric acid. 3. Dehydrate in acetone; mount. Results: Epidermis, yellow. Cortex, yellow. Pericycle (lignified tissues), blood red. Zylem, salmon to blood red. Primary phloem, orange. Cambium, pale yellow. Secondary phloem, crimson. Sieve plates, some very bright crimson, others orange. Pith, amber. Abstract: (1947) Stain Tech., VoL 22, 4, 155-6, B. Vercourt. COTTON BLUE - SAFRANIN For fungal h3rphae in woody tissues Solutions required: A. Cotton blue 0-5% in Lactophenol. B. Lactophenol. C. Safranin 1% aqueous. Technique: 1 . Stain thin sections with solution A, which has been warmed to about 35° C, for five to fifteen minutes. 2. Wash with Lactophenol to remove excess stain. 3. Wash with 70% alcohol to remove Lactophenol. 4. Counterstain with the safranin for about ten minutes. 5. Wash with 70% alcohol taking care that the safranin is not entirely removed. 6. Rinse rapidly in 90% alcohol. 7. Dehydrate with absolute alcohol. 8. Clear in xylol. 9. Mount in Canada balsam in xylol or in Cristalite. Results: Fungal hyphae, blue. Xylem, red. Reference: Chesters (1934), Ann. Bot., 48, 820. SECTION THREE CYANIN - ERYTHROSIN A botanical stain for cellulose and lignified tissues Solutions required: A. Cyanin, o*i% in 50% alcohol. Note: This is a very costly stain. B. Erythrosin 1% in 70% alcohol. Technique: 1. Stain sections from five to twenty minutes in the cyanin solution, placing the slide under a petri dish lid, or an evaporating basin lined with a piece of damped filter paper to prevent evapora- tion of the stain. 2. Wash rapidly with 50% alcohol. 3. Stain from half to one minute in the erjrthrosin. 4. Rinse quickly in 50%. 5. Rinse quickly in 95% alcohol. 6. Dehydrate by rinsing quickly in two changes of absolute alcohol. 7. Clear in xylol, and mount. Results: Lignified tissues are stained blue, while cellulose tissues are red. DELAFIELD HAEMATOXYLIN - CELLOSOLVE For botanical tissues, particularly for cell walls Solutions required: A. Delafield haematoxylin. B. Alcohol 70% . . . . . . 99*5 ml. Ammonia solution (sp. gr. o-88o) 0-5 cc. Technique: I. Fix sections to slides and stain in Delafield haematoxylin for three to five minutes. 255 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 2. Rinse in tap water. 3. Decolorize for about thirty seconds in 70% alcohol. 4. Rinse in 70% alcohol. 5. Blue in Solution B (above). 6. Rinse well in distilled water. 7. Immerse in two changes of cellosolve for one minute in each. 8. Drain and mount in Canada balsam or in Cristalite. I Results: Nuclei and chloroplasts are stained bluish purple; cytoplasm in a paler shade of purplish blue, or colourless, lignified and cutin- ized tissues are yellow. Unlignified tissues, bluish purple. ERYTHROSIN - LACTOPHENOL A general stain for botanical tissues Solution required: Erythrosin 1% in lactophenol. Technique: 1. Place sections on slides and add a drop of the staining solution. 2. Cover with a coverslip and examine under the microscope. Results: Lignified and cutinized tissues are stained yellow while cellulose walls are unstained. Nuclei and dense protoplasmic contents are red. Cytoplasm of vacuolated cells appear pink, while chloro- plasts, slime plugs in sieve, tubes, etc., are red. GRAM'S IODINE A general stain for botanical tissues Solutions required: A. Gram's iodine. B. Glycerine 50% aqueous. 256 SECTION THREE Technique: 1. Stain with Gram's iodine solution for a few minutes. 2. Pour off excess stain ; rinse in distilled water. 3. Mount in the glycerine solution; cover with a coverslip and examine under the microscope. Results: Starch, navy blue. Proteins, brown. Cytoplasm, light brown. Nuclei, dark brown. Chloroplasts, brown or blue. Cellulose walls, faint yellow. Lignified walls, deep yellow. HAEMATOXYLIN (Heidenhain) - ANILINE BLUE For the difTerential staining of nuclei, cytoplasm and cell walls of angiosperm shoot apices This technique which is due to Dr. J. G. Vaughan, Department of Biology, Chelsea Polytechnic, London, S.W.3, has been used with success on Anagallis and certain members of the Cruciferae. No mixing of the dyes has been observed in the apical meristem region, as occurs with other stain combinations, and the picture is clear and well defined, thereby facilitating study and photographing. Solutions required: A. Haematoxylin (Heidenhain), No. i. B. Haematoxylin (Heidenhain) No. 2. C. Iron alum 2% aqueous. D. Aniline blue alcohol, soluble (Michrome brand) . . . . . . . . i gm. Methyl cellosolve . . . . . . 100 ml. Dissolve by heating on a hot plate or a waterbath, taking care that the cellosolve does not catch fire. Allow to cool; then filter. E. Methyl salicylate 25 ml. Xylol Absolute alcohol F. Methyl salicylate Xylol Absolute alcohol 33 nal- 42 ml. 40 ml. 20 ml. 20 ml. 257 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES G. Xylol . . . . . . . . 90 ml. Absolute alcohol . . . . . . 10 ml. Technique: 1. Pass sections through descending grades of alcohol to dis- tilled water. 2. Mordant in solution A for thirty minutes. 3. Rinse in distilled water. 4. Stain in solution B for twelve hours. 5. Rinse in distilled water. 6. Differentiate in 2% iron alum solution. 7. Wash in running water for one hour. 8. Dehydrate in 25%, 50%, 70%, 90% and two changes of absolute alcohol. 9. Stain in the aniline blue solution for ten minutes. 10. Remove excess aniline blue with absolute alcohol. 11. Rinse in solution E. 12. Clear in solution F for ten minutes. 13. Rinse in solution G. 14. Rinse in two changes of xylol and mount in Canada balsam in xylol, Clearmount, Cristalite or D.P.X. Results: Nuclei and cytoplasm are well stained by the haematoxylin, while cell walls are stained very satisfactorily by the aniline blue. Note: (a) The success or failure of the method is said to depend on the quality of the aniline blue. - [h) Ilford Special Rapid Panchromatic plates were used for the preparation of the photomicrographs which are reproduced in the original paper. Reference: Vaughan, J. G. (1955), Stain Tech., 30, no. 2, 79-82. 258 SECTION THREE HAEMATOXYLIN - BISMARK BROWN For Phloem tissues of woody plants Solutions required: A. Iron alum 2% aqueous. B. Haematoxylin i% aqueous. C. Bismark Brown 1% aqueous. Technique: 1. Immerse sections in solution A for ten to twenty minutes. 2. Drain off excess solution; then wash in six or seven changes of distilled water. 3. Cover the preparation with solution B, placing the slide on the microscope stage. 4. Observe the progress of the stain under the low power objective and when the required depth of staining has been at- tained, pour off excess stain and rinse quickly in distilled water. 5. Rinse again with two or three changes of distilled water. 6. Immerse the preparation in solution C in a staining tube or jar for three to four hours, depending on the thickness of the sections. 7. Remove excess stain by washing well with water. 8. Wash with 50% alcohol, followed by 70% alcohol. 9. Wash with 90% alcohol. 10. Wash with two changes of absolute alcohol. 11. Clear in xylol, and mount in Cristalite or Clearmount or Canada balsam in xylol. Results: Stone cells of hard woods are stained cherry red; bast fibres are brilliant orange, whilst the ray cells and other parenchymatous tissues are a chestnut brown, and the middle lamellae dark blue. The bast fibres, parenchymatous tissues, and middle lamellae of Coniferophyta are stained as indicated, but the stone cells turn a vivid burnt orange. 259 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES HEroENHAIN HAEMATOXYLIN - SAFRANIN A general stain for plant tissue, algae, fungi, etc., to demon- strate histological and cytological structures Solutions required: A. Haematoxylin (Heidenhain) No. i. B. Haematoxylin (Heidenhain) No. 2. C. Picric acid, saturated, aqueous. D. Safranin 1% aqueous. Technique: 1 . Fix and dehydrate tissues and embed in paraffin wax. Note: Algae and fungi should be treated by the Venetian tur- pentine method {see page 43). 2. Fix paraffin sections to slides ; remove wax with xylol. 3. Pass through absolute alcohol and the usual descending grades of alcohol, down to distilled water. 4. Mordant in Heidenhain haematoxylin No. i (Solution A) from one half to two hours but no longer, unless the material is algae in which case as long as twelve hours may be necessary. 5. Wash for five minutes in running tap water. 6. Rinse well in distilled water. 7. Stain algae for at least twenty-four hours, or other material from five to seven hours, in Heidenhain haematoxylin No. 2 (Solution B). 8. Wash in running water for five minutes. 9. Differentiate in Solution C (picric acid) from twenty minutes to two hours, controlling by examination at intervals under the microscope. 10. Wash for half an hour in running tap water. 11. Rinse well in distilled water. 12. Counterstain in the safranin solution for five to ten minutes. 13. Rinse in distilled water. 260 SECTION THREE 14. Dehydrate as usual. 15. Immerse for five minutes in equal parts of absolute alcohol and xylol. 16. Transfer to and immerse in xylol for five minutes. 17. Mount in Canada balsam in xylol. Results: Chromosomes, black to purple. Centrosomes and pyrenoids, black to purple. Lignified, suberized and cutinized structures, un- stained or only faintly stained. Archesporial cells and early stages of sporogenous tissue, grey. IODINE GREEN - ACID FUCHSIN A botanical stain for lignified tissues, and for chromosomes Solutions required: A. Iodine 1% aqueous. B. Acid fuchsin 1% aqueous. Technique: 1 . Fix material in Nevashin's fluid. 2. Take sections through to distilled water in the usual manner. 3. Immerse for at least twelve hours in the Iodine green solution. 4. Wash and differentiate with distilled water, examining at intervals under the microscope while the preparation is still wet, until the non-lignified tissues retain only a faint green tint. 5. Counterstain for about fiYQ minutes in the acid fuchsin solution, controlling by examination under the microscope, while the preparation is still wet, taking care to ensure that the acid fuchsin solution is not allowed to act long enough to extract the Iodine green from the lignified tissues. 6. Rinse quickly in 90% alcohol. 7. Dehydrate quickly in two changes of absolute alcohol. 8. Clear in clove oil. 9. Wash with xylol. 261 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 10. Mount in Cristalite, Clearmount or in Canada balsam in xylol. Results: Chromosomes and nuclei, green. Plastin and cytoplasm, pink to red. Lignified tissues, green. JOHANSEN'S QUADRUPLE STAIN For plant tissue Solutions required: A. Safranin 0, i% in cellosolve 50 ml. Alcohol 95% 25 ml. Distilled water 25 ml. Sodium acetate . . I gm. Formalin . . 2 ml. B. Crystal violet i% aqueous. C. Alcohol 95% 25 ml. Cellosolve 25 ml. Tertiary butyl alcohol 25 ml. D. Fast Green, FCF, saturated in equal parts of clove oil and cellosolve 20 ml. Alcohol 95% 60 ml. Tertiary butyl alcohol . . 60 ml. Glacial acetic acid 0-6 ml. E. Alcohol 95% 30 ml. Tertiary butyl alcohol 30 ml. Glacial acetic acid 0-15 ml F. Orange G saturated in cellosolve 20 ml. Cellosolve 20 ml. Alcohol 95% 20 ml. G. Clove oil . . 10 ml. Absolute alcohol . . 10 ml. Xylol 10 ml. 262 SECTION THREE Technique: 1. Paraffin sections are brought down to 70% alcohol; then stained for twenty-four to forty-eight hours in Solution A (over- staining is not possible). 2. Rinse in water ; then stain ten to fifteen minutes in Solution B. 3. Rinse in water; then rinse for fifteen seconds in Solution C. 4. Stain ten to twenty-five minutes, according to the material and fixative, in Solution D. 5. Rinse briefly in Solution E. 6. Immerse for about three minutes in Solution F. 7. Rinse in Solution G. 8. Rinse in two changes of xylol ; then mount in balsam. Results: Dividing chromatin, red ; resting chromatin, purplish ; nucleoli, red (occasionally violet); nucleoplasm, colourless or greenish; lignified walls, bright red; cutinized cell walls, reddish purple; suberized walls, red ; cellulose cell walls, greenish orange ; cyto- plasm, bright orange ; middle lamellae, green ; starch grains, purple with green or orange halos (the colour of the halps soon becomes replaced by purple in some types of materials) ; plastids, purplish to greenish; invading fungal mycelium, green; the callose portion of the guard cells of the stromata bright red and the remainder purple ; and Casparian strips, red ; the remainder of the cell wall of the endodermis, yellow. In sections of roots for the origin of the lateral roots, the cytoplasm of the latter should be stained green, with purplish nuclei, while the cytoplasm elsewhere should be orange with red nuclei. The combination is exceptionally good for sections of lichens, as the algae are well differentiated, and also for Puccinia graminis telia and Uredinia. From Plant Microtechnique, by D. A. Johansen, by courtesy of McGraw-Hill Book Company, Inc., New York. T 263 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES JOHANSEN'S QUINTUPLE STAIN For plant tissue composed of a variety of cell types, such as leaves, roots, steins and ovaries Solutions required: A. Safranin O, i% in cellosolve . . 50 ml. Alcohol 95% . . . . . . 25 ml. Sodium acetate 4% . . . . 25 ml. Formalin . . . . . . . . i ml. B. Crystal violet 1% aqueous. C. Absolute alcohol Cellosolve Tertiary butyl alcohol D. Fast Green, FCF. . Malachite Green . . Cellosolve Dissolve; then add: Tertiary butyl alcohol Absolute alcohol Glacial acetic acid E. Orange 2 . . Cellosolve Dissolve by warming gently, Clove oil . . Absolute alcohol Tertiary butyl alcohol F. Terpineol, extra pure Absolute alcohol Tertiary butyl alcohol . . I equal Cellosolve . . . . f volumes Methyl salicylate Beechwood creosote 25 ml. 25 ml. 25 ml. 0-5 gm. 0-5 gm. 100 ml. 25 ml. 25 ml. 1*5 ml. 0-5 gm. 58 ml. then add : 14 ml. 14 ml. 14 ml. 264 SECTION THREE G. Terpineol, extra pure Absolute alcohol Tertiary butyl alcohol . . Cellosolve Methyl salicylate Beechv/ood creosote i equal volumes Toluol H. Toluol . 45 ml. Absolute alcohol . 5 ml. Technique: 1. Paraffin sections are fixed to slides, and taken down through the usual stages to 70% alcohol. 2. Stain for four to forty-eight hours in the safranin (Solution A). Note: The staining time depends upon the material. Angio- sperm and Pteridophyte stems, leaves and roots, usually require at least twenty-four hours. Weakly lignified or overchromated tissues may need forty-eight hours. Gymnosperm materials need any- thing from four to forty-eight hours. Ovaries and embryos require four hours. 3. Rinse thoroughly in water. 4. Stain for five minutes in the crystal violet (Solution B). Note: This stain should be omitted on ovules, embryos and all parasitic and saprophytic fungi ; in such cases proceed from stage 3 directly to stage 7. 5. Rinse in running water for a few seconds. 6. Rinse for a minute in Solution C. 7. Stain for five minutes in the fast green - malachite green (Solution D). 8. Rinse briefly in running water. Note: The preparations will appear to be excessively over- stained, but this does not matter. 9. Rinse in Solution C to which 1% acetic acid has been added. 10. Stain for five minutes in the orange 2 (Solution E). 1 1 . Rinse in Solution F for a minute. 265 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 12. Rinse in Solution G. 13. Immerse in Solution H for five to ten minutes. 14. Pass through two changes of alcohol; then mount. Notes: {a) Isopropyl alcohol may be used, if desired, for every purpose for which ethyl alcohol is commonly used. {h) The schedule at first glance appears to be complicated, but the actual application is quite simple. {c) Slides should be agitated in all rinses and washes. (d) The reason for mixing so many reagents together in Solu- tions F and G is that each effects certain stains and not the others. {e) The fast green - malachite green (Solution D) is somewhat strong at first but gradually weakens as successive slides are passed through it ; consequently, the time in this stage may be reduced to four minutes and that of the orange 2 (Solution E) in Stage 10 leng- thened to ten minutes, then equalized, and later, the changes in time reversed as the orange becomes stronger after its initial weak- ness. ( / ) Approximately three hundred slides may be passed through each 100 ml. of each staining solution (except the safranin (Solu- tion A), which merely needs replenishment when required), and rinsed before replacement becomes necessary owing to contamin- ation. From personal communications with Professor D. A. Johansen, Pomona, California, U.S.A., to whom this technique and my thanks are due. LACMOID - TANNIC ACID - FERRIC CHLORIDE For phloem and contiguous tissues: the technique gives relatively stable preparations of critically stained materials Solution required: A. Tannic acid 1% aqueous. B. Ferric chloride, hydrated 2% aqueous. C. Sodium bicarbonate . . . . 2-5 gm. Distilled water . . . . • . 50 ml. Dissolve by shaking or stirring: do not apply heat, 266 SECTION THREE D. Solution C . . Distilled water Absolute alcohol E. Lacmoid Absolute alcohol Distilled water Solution C . . F. Solution C . . Distilled water Absolute alcohol G. Absolute alcohol Clove oil Xylene 20 ml. 55 ml. 25 ml. 0-25 gm. 30 ml. 70 ml. 3 to 5 ml. 20 ml. 30 ml. 50 ml. equal volumes of each Technique: 1. Fix material in formalin-acetic-alcohol. 2. Unembedded sections, 5 to 40/x in thickness may be em- ployed. 3. Sections are taken from distilled water and immersed in 1% tannic acid for five to ten minutes. 4. Transfer to 2% ferric chloride for about five mins. 5. Wash in distilled water (three changes). 6. Examine under the microscope while the preparation is still wet : the colour of the walls should be medium to dark grey, and it may be necessary to repeat the above staining process to attain this result as some phloem tissues stain less readily than others. 7. Wash in three changes of distilled water. 8. Immerse in solution D for thirty minutes. 9. Transfer directly to solution E and leave therein for twelve to eighteen hours, or longer if desired, as it is impossible to over- stain. 10. Transfer to solution F from ten seconds to ten minutes depending upon the time at which the lacmoid destains. 1 1 . Wash in 80% alcohol. 12. Immerse in 90% alcohol for two to three minutes. 267 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 13. Immerse in two changes of absolute alcohol for a total time of two to three minutes. 14. Wash in solution G for two to three minutes. 15. Immerse in two changes of xylol for a total time of two to three minutes. 16. Mount in D.P.X. or Clearmount or Cristalite mountant. Results: Callose is stained sky blue to greenish blue; lignified cellulose (in the xylem cells and in most cortical or phloem fibres and sclereids) blue. Cellulose walls, nuclei, slime and cytoplasm, light brown to greyish brown. Reference: Vernon, J. C. and Gifford, E. M. (1953), Stain Tech., 28, 49-53. LACMOID - MARTIUS YELLOW For callose in pollen tubes Solution required: Lacmoid 0-01% .. .. .. 10 ml. Martins Yellow 0-01% .. .. lo ml. Add a few drops of diluted ammonia (o-i ml. cone, ammonia with 10 ml. distilled water) until the solution assumes an olive tint. Technique: 1. Slender styles or ovaries are crushed between two slides while still wet. Larger styles or ovaries should be cut by hand into longi- tudinal sections, then crushed. 2. Stain two to five minutes in the Lacmoid - Martins yellow solution. 3. Mount in the stain and examine under the microscope, using a strong light. Results: Pollen tubes, blue. Background, light yellowish green. 268 SECTION THREE LIGNIN PINK A non-fading stain which is specific for lignin The stain gives constant results with reasonably thin sections and is particularly suitable for routine or elementary classwork. Overstaining with Lignin Pink is impossible, and it will not wash out with alcohol. In combination with chlorazol black it offers a simple double staining technique as follows : Solutions required: Lignin Pink . . Chlorazol black Distilled water • 0-5 gm. , . 0-5 gm. . 100 ml. Technique: 1. Reasonably thin sections are fixed to slides and immersed in the staining solution for twenty to twenty-five minutes. 2. Rinse in distilled water. 3. Rinse in 70% alcohol. 4. Rinse in 90% alcohol. 5. Dehydrate with absolute alcohol. 6. Clear in xylol and mount. Results: Lignin stands out, stained bright carmine colour, against the surrounding tissues which are stained black. Reference: Cannon, H. G. (1941)^^. jR. Mic. Soc, series III, 61, parts 3 and 4. MAGDALA RED - FAST GREEN A differential stain for parasite and host tissues in botanical material Solutions required: A. Magdala red 2% in 80% alcohol (original) B. Fast Green FCF-Clove Oil. 269 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Embed the material in paraffin wax in the usual manner. 2. Remove paraffin wax from the sections with xylol as usual. 3. Wash well with absolute alcohol. 4. Wash with 90% alcohol. 5. Stain for five to ten minutes with solution A. 6. Wash with absolute alcohol to remove excess stain. 7. Counterstain for two to five minutes in solution B. 8. Wash in absolute alcohol. 9. Clear in xylol, and mount. Results: Host tissues are stained green ; parasite, red. METHYL GREEN - PHLOXIN - GLYCERINE JELLY A rapid method for the simultaneous mounting and double staining of pollen grains, differentiating functional from abortive grains This technique can be employed for the evaluation of orchid seeds which contain naked embryo surrounded by integuments. Solutions required: A. Methyl Green, saturated in 50% alcohol. B. Phloxin, saturated in 50% alcohol. C. Glycerine jelly . . . . . . 50 ml. Solution A . . . . . . . . 2J ml. Solution B . . . . . . . . 2 ml. Melt glycerine jelly on a water bath ; then measure off in a pre- heated measuring cylinder the 50 ml. required and pour this amount into a pre-heated bottle to which, solutions A and B are then added and shaken in. Note: The jelly should now be a port wine colour: it may he necessary to vary the proportions by adding more of one of the dyes to produce this colour. 270 SECTION THREE Technique: 1. Place a small amount of pollen on the centre of a slide. 2. Wash by dropping on 70% alcohol to remove any adhering oils and resins. Note: The alcohol will spread out and evaporate leaving a ring of sludge, which should he removed with a piece of tissue paper moistened with alcohol. 3. Repeat this process until no more sludge comes out. 4. Wash finally with 70% alcohol, and just before the last drop of alcohol evaporates completely, place 2 drops of hot stained glycerine jelly on the pollen and stir gently with a needle to ensure even distribution. 5. Cover with a coverslip, keeping the jelly under the coverslip hot by heating gently with a bunsen flame. Note: Excessive heating will rupture many of the pollen grains. 6. Remove the flame ; take off the coverslip and replace it with a clean, flamed coverslip. 7. Allow the sUde to cool ; then store in a cool place. Results: ''Functional" pollen grains are fully expanded; exine and intine are stained green ; cytoplasm red. Aborted grains are either shrunken and stained green, or expanded in varying degrees depending on the amount of non-autolysed cytoplasm present at the time of mounting, and stained a mottled, reticulate red. Notes: The intensity of the Phloxin stain increases on standing, whereas the action of the methyl green is practically instantaneous. Preparations retain their brilliancy for about a year. Reference: Owczarzak, Alfred (1952), Stain Tech., 27, no. 5, p. 249. PHLOROGLUCINOL An extremely sensitive test for lignin, particularly suitable for hydrophytes Solution required: Phloroglucinol 1% in 70% alcohol. 271 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Sections are placed on slides and flooded with the phloro- glucinol solution which is allowed to act for about five minutes. 2. Pour off excess phloroglucinol and cover sections with a few drops of concentrated hydrochloric acid, cover with a coverslip and examine under the microscope. Result: Lignified tissues are stained red. POLYVINYL LACTOPHENOL For embedding brittle specimens of wood for sectioning. It is claimed that this technique has given successful results with wood dating back to the Roman era In addition to softening the wood for cutting, this technique also clears the specimen and allows cell walls, which may have been blackened through carbonization, to become clear and translucent, thereby facilitating identification under the microscope. Solution required: Polyvinyl Lactophenol. Technique: 1. Immerse blocks of wood cubed in the usual manner, in polyvinyl alcohol, and warm gently for thirty minutes. 2. Drain, and allow to cool for twenty-four hours. 3. Cut sections from blocks with a very sharp razor. Results: The wood has lost its brittleness and has acquired a soft pliable rubber-like nature. Reference: Levy, J. F. L. (1953), Nature, 171, 984. 272 SECTION THREE SAFRANIN - ANILINE BLUE For plant tissues; particularly suitable for gymnosperm ovules, archegonia, embryos and angiosperm stems and roots Solutions required: A. Safranin O, 2% in Cellosolve . . 100 ml. Absolute alcohol . . . . . . 50 ml. Sodium acetate 4% aqueous . . 50 ml. Formaldehyde 40% . . . . . . 8 ml. B. Picric acid 0*5% in 95% alcohol. C. Ammonia solution (sp. gr. 0.88) . . 0-25 ml. Absolute alcohol . . . . . . 100 ml. D. Aniline Blue, alcohol soluble, saturated in equal volumes of cellosolve and absolute alcohol. Technique: 1. Take sections through to 70% alcohol in the usual manner. 2. Stain from two to forty-eight hours, according to the nature of the material, in solution A. Note: Gymnosperms require the minimum staining time, 3. Wash thoroughly with running water to remove the excess stain. 4. Dehydrate and differentiate carefully with solution B. 5. Immerse the preparation in solution C for half to one minute. 6. Dehydrate morphological material in 95% alcohol, or cyto- logical preparations in absolute alcohol. 7. Counterstain for about one minute in a mixture consisting of equal parts of solution D and clove oil. 8. Clear in methyl salicylate and mount in Cristalite, Canada balsam or Emexel mountant. Results: Lignified and cutinized cell walls, nuclei and chromosomes, bright red. Cellulose cell walls and cytoplasm, blue. 273 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Note: At certain developmental and formative stages, some cell walls are stained sharply with the safranin, while other portions will appear faint blue. SAFRANIN - ANILINE BLUE For plant tissues, particularly for chromosomes and cell walls Solutions required: A. Safranin i% aqueous. B. Aniline blue i% in absolute alcohol. Technique: 1 . Paraffin sections are fixed to slides ; de- waxed with xylol, and passed through descending grades of alcohol down to distilled water as usual ; alternatively freehand sections may be employed. 2. Stain for fifteen minutes in the safranin solution. 3. Wash in distilled water. 4. Rinse in 70% alcohol, followed by 90%. 5. Rinse in absolute alcohol. 6. Stain for two minutes in the aniline blue solution. 7. Rinse quickly but thoroughly with two changes of absolute alcohol. 8. Wash with two changes of xylol. 9. Mount in balsam. Results: Chromosomes, red. Nucleoli, red. Cytoplasm almost colourless. Cellulose walls, blue. SAFRANIN - DIANIL BLUE G For the differential staining of Peronosporaceae Solutions required: A. Phenol crystals . . . . . . 10 gm. Lactic acid . . . . . . . . 10 ml. Glycerin . . . . . . . . 20 ml. Absolute alcohol . . . . . . 20 ml. 274 SECTION THREE B. Cotton blue 4B . . . . . . 0-02 gm. Safranin . . . . . . . . o-io gm. Solution A . . . . . . . . 100 ml. C. Safranin 0*25% in clove oil,. Technique: 1. Embed material in paraffin wax by the standard technique. 2. Fix sections to slides and remove paraffin wax with xylol as usual. 3. Wash well with absolute, followed by 90% alcohol. 4. Immerse in solution A for ten to fifteen minutes. 5. Transfer to solution B and leave therein for two hours. 6. Differentiate in solution A. 7. Wash with absolute alcohol. 8. Immerse in solution C for twenty to thirty minutes. 9. Differentiate in clove oil. 10. Clear in xylol, and mount in Canada balsam in xylol or in Cristalite, or in Emexel mountant. Results: Host tissue is stained red ; Myelium blue. Reference: (1928), Lepik. Phtopath., 18, 869. SAFRANIN - FAST GREEN IN CELLOSOLVE A rapid, non-fading stain for botanical tissues in place of Safranin -light green -clove oil Solution required: 2 % Safranin - Fast Green FCF in cellosolve. Technique: 1. Stain sections for five to ten minutes. 2. Rinse in cellosolve. 4. Mount in Canada balsam or in CristaHte. 275 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Results: Lignified tissues are stained red, while unlignified tissues and cytoplasm are blue-green. Nuclei, red. Chloroplasts, pink to red. SAFRANIN - ACID FUCHSIN For spermatozoids, zoospores, motile gametes, etc. Solutions required: A. Osmic acid i% aqueous. B. Safranin i% aqueous. C. Acid Fuchsin i% aqueous. Technique: 1 . Place a drop of an aqueous suspension of the organism on a slide. 2. Fix by holding the slide for about one minute over osmic acid solution. 3. Allow the preparation to dry. 4. Stain for ten minutes to one hour in solution B. 5. Wash in water. 6. Remove excess water by draining and blotting the edges of the slide with filter paper. 7. Wash in 95% alcohol until only the nuclei retain the stain. 8. Stain in the acid fuchsin solution for ten to twenty seconds. 9. Wash rapidly in 70% followed by 90% alcohol. 10. Wash rapidly with two changes of absolute alcohol. 11. Clear in clove oil, followed by xylol. 12. Mount in Canada balsam in xylol, Cristalite, Clearmount or Emexel mountant. Results: Nuclei bright red. Cytoplasm bluish pink. Reference: (19 18), Steil. Bot. Gaz., LVX, 592. 276 SECTION THREE SAFRANIN - FAST GREEN, FCF A non-fading stain, satisfactory for nearly every type of plant material except the algae Solutions required: A. Safranin O, alcoholic . . I gm. Cellosolve . . 50 ml Alcohol 95% . . 25 ml Distilled water . . 25 ml Sodium acetate . . . . I gm. Formalin . . . . 2 ml. B. Fast Green FCF: Fast Green FCF saturated in equal parts Cellosolve and absolute alcohol added in sufficient quantity to give a stain of the desired intensity when mixed with a mixture consisting of one part absolute alcohol and three parts clove oil. t Technique: 1. Paraffin sections are brought down to 70% alcohol, or free- hand sections up to 35% alcohol. 2. Stain in Solution A for two to forty-eight hours (gymno- sperm material needs the minimum period) ; then wash off excess stain in running water for a minute or so. 3. Differentiate and dehydrate simultaneously with 0-5% picric acid in 95% alcohol. 4. Immerse in 95% alcohol to which four or five drops of ammonia have been added from a few seconds to two minutes, but no longer. 5. Dehydrate quickly with absolute alcohol. 6. Counterstain with Solution B fdr ten to fifteen seconds. 7. Wash off excess stain with clove oil then clear in a mixture consisting of two parts of clove oil, one part absolute alcohol, and one part xylol. 8. Remove the clearing mixture by washing for a few seconds each in three changes of xylol ; then mount in balsam. 277 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Results: Nuclei, chromosomes, lignified and cutinized cell walls, brilliant red. Cytoplasm and cellulose cell wall, brilliant green. In some cell walls at certain developmental or formative stages, portions will be stained less sharply by safranin, and other portions weakly by the fast green. SAFRANIN - FAST GREEN, FCF IN CELLOSOLVE A rapid non-fading stain for botanical tissues in place of Safranin - light green - clove oil Solution required: Safranin - light green in Cellosolve. Technique: 1. Stain sections in the safranin - light green - cellosolve for five to ten minutes. 2. Rinse in Cellosolve. 3. Mount in Canada balsam or in Cristalite. Results: Lignified tissues are stained red, while unlignified tissues and cytoplasm are blue-green. Nuclei, red. Chloroplasts, pink to red. SAFRANIN - LIGHT GREEN - CLOVE OIL A general stain for botanical tissues Solutions required: A. Safranin 1% in 50% alcohol. B. Light Green - clove oil. Technique: 1. Stain section in the safranin solution for ten minutes. 2. Rinse in two changes of 50% alcohol for thirty seconds in each. 278 i SECTION THREE 3. Rinse in two changes of 70% alcohol for thirty seconds in each. 4. Repeat with two changes of 90% alcohol for the same time. 5. Immerse in two changes of absolute alcohol for two or three minutes in each. 6. Stain in light green - clove oil for one minute. 7. Rinse and wash in clove oil for about five minutes. 8. Examine under the microscope and if it is found that the safranin has been extracted by the alcohols, take the section down through the alcohols and restain with safranin. 9. Mount in balsam or in Cristalite mountant. Results: Cellulose tissues and cytoplasm are stained green. Lignified tissues and nuclei, red. Chloroplasts, pink. SAFRANIN - PICRO ANILINE BLUE A rapid and simple method of demonstrating hyphae in wood sections Solutions required: A. Safranin 1% in distilled water. B. Aniline blue, water soluble . . 2-5 gm. Distilled water . . . . . . 25 ml. Picric acid, saturated, aqueous . . 100 ml. Technique: 1. Stain one to three minutes in Solution A ; then wash in water. 2. Flood sections with Solution B and heat over a flame until it begins to simmer ; then pour oif excess stain ; allow the slide to cool before washing with water. 3. Wash with 70%, followed by absolute, alcohol ; clear in clove oil ; mount in balsam. Results: Lignified walls, red. Mycelia, clear blue. Areas where the wood is badly decayed may appear bluish, but the hyphae are always well defined. u 279 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES SAFRANIN - TANNIC ACID - FAST GREEN For roots and steins Solutions required: A. Tannic acid i% in 50% alcohol. B. Ferric chloride 3% in 50% alcohol. C. Safranin 1% in 50% alcohol. D. Fast green FCF in clove oil to which has been added 7% of absolute alcohol. Technique: 1. Sections are fixed to slides, treated with 70% alcohol; then immersed in the tannic acid solution. 2. Immerse for about twenty seconds each in two lots of 50% alcohol. 3. Treat with the ferric chloride for about thirty to sixty seconds. 4. Wash with 50% alcohol. 5. Stain with the safranin solution for twenty-four hours, in a stoppered staining jar. 6. Wash with 50% alcohol. 7. Differentiate for about ten seconds in 70% alcohol. 8. Pass through 80%, 90% and absolute alcohol, allowing ten to thirty seconds in each. 9. Stain for two or three minutes in the fast green. 10. Pass through absolute alcohol, followed by xylol; then mount in Cristalite. Results: Nuclei, red. Cytoplasm, blue-green. Lignified walls, red. Cambial cell walls, black. Collenchyma walls, very dark red. Parenchyma cell walls, black. 280 SECTION THREE SCARLET R or SUDAN 3 - ETHYLENE GLYCOL An improved technique for staining fat in plant tissues, offering the following advantages : (a) The use of ethyl alcohol is obviated and sections remain pliable and unshrunken; {b) excellent dif- ferentiation without loss of stain from the fat; (c) More intense staining of fat. Solutions required: A. Scarlet R or Sudan 3 . . . . i gm. Ethylene glycol, pure, anhydrous.. 100 ml. Heat the ethylene glycol to 100-110° C. on a hot plate, or in an oven for preference, but if these are not available the bunsen flame will serve the purpose so long as care is taken to ensure that the ethylene glycol does not take fire. Stir in the dye until all or most of it is dissolved; then cool and filter. B. Ethylene glycol . . . . • • 85 ml. Distilled water . . . . . . 15 ml. C. Delafield or Ehrlich haematoxylin. Technique: 1. Fix tissues in 10% formalin. 2. Wash out fixative with water. 3. Dehydrate the sections by agitating gently in the pure ethylene glycol, anhydrous, for three to five minutes. 4. Immerse the sections in the staining solution (Scarlet R or Sudan 3) and agitate for two to five minutes. 5. Differentiate by agitating gently, at intervals, from one to five minutes in 85% ethylene glycol (solution B) controlling by examination under the microscope while the specimen is still wet. 6. Transfer the sections to distilled water and leave therein for three to five minutes. 7. Counterstain with Delafield or Ehrlich haematoxylin. 8. Wash well in tap water. 9. Mount in glycerine jelly or glycerine. 281 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Results: , With Scarlet R: Nuclei, blue. Fat, orange to red. Cholesterol, j red. Fatty acids, unstained. i With Sudan 3 .* Nuclei, blue. Fat, yellow to orange. Cholesterol, \ orange. Fatty acids, unstained. SCARLET R For staining fat in plant tissues Solutions required: ^ A. Scarlet R, saturated in 70% alcohol. j B. Delafield or Ehrlich haematoxylin. Technique: 1. Fix tissues in 10% formalin. 2. Sections are immersed for a second in 70% alcohol; then stained for two to five minutes in the Scarlet R. 3. Wash quickly and transfer to distilled water. 4. Counterstain with Ehrlich or Delafield haematoxylin. 5. Wash well in tap water; mount in glycerine or glycerine jelly. Results: Fat, orange to red. Cholesterol, red. Nuclei, greyish blue. Fatty acids, unstained. SCHULZE SOLUTION (Chlor Zinc Iodine) For cell walls, proteins and starch 1. Place sections on slides and pour on a few drops of Schulze solution. 2. Cover with a coverslip and examine under the microscope. Results: Starch, blue. Proteins, brown. Cellulose walls, violet. Ligni- fied walls, yellow. Cutinized and suberized walls, yellow to brown. 282 SECTION THREE TETRAZOLIUM SALT For testing the viability of seeds Solution required: A. Tetrazolium salt i% aqueous. Technique: 1. Select the seeds at random and steep in tap water for i8 hours. 2. Section longitudinally through the embryo and place one half of each in a 7 cm. petri dish. 3. Pour the tetrazolium salt solution over the seeds and soak in the dark for 4 hours at 20° C. 4. Wash with tap water. 5. Examine seeds for staining. Results: Viable seeds are stained: others unstained. Note: The method is not well adapted to certain seeds. Reference: Cottrell, Helen J. (1948), Ann. Appl. Biol., 35, 123-31. THIONIN - ORANGE G For infected plant tissues Solutions required: A. Carbol thionin. B. Orange G 0-2% in absolute alcohol. Technique: 1. Fix paraffin sections to slides; pass through xylol and descending grades of alcohol down to distilled water in the usual manner. 2. Stain for an hour in the thionin solution. Alternatively, free- hand sections may be employed, in which case, the sections should be stained in the thionin for only five minutes. 3. Pour off excess stain, and rinse with 70% alcohol followed by 90%. 4. Rinse quickly in two changes of absolute alcohol. 283 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 5. Differentiate for one half to one minute with the orange G solution. 6. Pass through 90%, followed by 70%, alcohol. 7. Rinse well with water. 8. Rinse with 70% alcohol, followed by 90%. 9. Rinse quickly in two changes of absolute alcohol. 10. Pass through two changes of xylol. 1 1 . Mount in balsam or in Cristalite. Results: Parasites are conspicuously stained violet-purple, with deep purple nuclei. Cell walls unstained. Note: Alternatively, stages 6, 7 and 8 may be deleted in which case the results will be as follows : parasites not so conspicuously stained violet-purple, with deep purple nuclei. Cellulose walls, yellow to green. Lignified tissue, blue. Tissue nuclei, pale blue with purple nucleoli, and chromosomes deep blue on a purple spindle. TITAN YELLOW For the detection of magnesium in plant cells Solutions required: A. Titan yellow, special for magnes- ium test 0*2% aqueous. B. Sodium hydroxide 10% aqueous. Technique: 1 . Paraffin sections are mounted on slides and brought down to distilled water in the usual manner, or freehand sections of fresh material may be employed. 2. Add one or two drops of the Titan yellow solution, followed by one or two drops of the sodium hydroxide solution. Results: If magnesium is present a red coloration is produced. 284 SECTION THREE TRYPAN BLUE A nuclear stain for plant material Solutions required: A. Formol Acetic alcohol ( Telly esniczky) Formaldehyde 40% 70% Alcohol Glacial acetic acid . . B. Trypan blue 1% aqueous . . Ethylene glycol Absolute alcohol Zinc sulphate C. Trypan blue 1% aqueous . . Distilled water Absolute alcohol Cresol HCl, concentrated . . D. Exactly the same as solution C but omit the HCl. E. Absolute alcohol . . . . . . 95 ml. Cresol . . . . . . . . 5 ml. F. Naphthol yellow, saturated in equal volumes of alcohol and ethylene glycol. G. Sodium thiocyanate 0-5% in equal volumes of ethylene glycol and absolute alcohol. H. Ponceau 3R satd. in 80% alcohol i volume 80% alcohol . . . . . . I volume 5 ml. . . 100 ml. 5 ml. . . 50 ml. . . 25 ml. . . 25 ml. . . 5 gm- . . 20 ml. . . 10 ml. . . 60 ml. . . 10 ml. . . o-i ml. Technique: 1. Fix epidermis or smears in solution A. 2. Stain epidermis overnight in solution B or C. Note: Anther smears are stained in solution D for one half to two hours. 3. Differentiate in 50% alcohol (epidermis for about fifteen minutes; smears for about five minutes), controlling by micro- 285 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES scopic examination, observing the nuclei rather than the non- nuclear material, as it may not be possible to remove all the blue stain from the latter without completely decolorizing the nuclei. Note: If 50% alcohol is found inadequate, then solutions E, F, and G should be tried in turn to ascertain which is the most satisfactory for the particular plant material. Some material, e.g. sweet clover epidermis needs little or no differentiation. Solutions F and G are more active as differentiators than E or 50% alcohol. 4. Place in 95% alcohol for two minutes. 5. Counterstain epidermis if desired with solution H for one minute but this may not be advantageous in the case of smears. 6. Rinse in 95% alcohol. 7. Immerse in two changes of Isopropyl alcohol for two minutes in each. 8. Immerse in a third change of Isopropyl alcohol for five minutes. 9. Clear in xylol and mount in Clearmount or balsam. Results: Nuclei, bright blue. If counterstain is used, the background is bright red. Reference: Hoffmesster, E. R. (1953), Stain Tech., 28, no. 6, 309. 286 SECTION 4-CYTOLOGICAL METHODS I ACID FUCHSIN - TOLUIDINE BLUE - AURANTIA For mitochondria Solutions required: A. Champy^s Fluid: Potassium dichromate 3% . . 7 ml. Chromic acid 1% . . . 7 ml. Osmic acid 2% . . . . 4 ml. B. Pyroligneous acid . . I volume Chromic acid 1% . . 2 volumes C. Potassium dichromate 3%. D. Acid fuchsin . . 10 gm. Aniline water . . 100 ml. E. Toluidine blue 0-5% aqueous. F. Aurantia 0-5% in 70% alcohol. Technique: 1. Tissues are fixed in Champy's Fluid for twenty-four hours; then washed in running water for at least an hour. 2. Immerse in Solution B for twelve to twenty hours ; then wash in distilled water for thirty minutes. 3. Mordant in Solution C for three days; then wash in running water for twenty-four hours. 4. Dehydrate ; clear ; embed in paraffin wax. 5. Sections, 3 to 5yW in thickness are taken down to water in the usual manner ; then stained for six minutes by flooding the slide with Solution D, and heating gently till vapour rises. 6. Rinse in distilled water; then counterstain in Solution E. 289 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 7. Rinse with distilled water; then stain with Solution F for thirty to fifty seconds. 8. Differentiate with 95% alcohol; dehydrate; clear in xylol and mount. Results: Mitochondria are stained red; nuclei, blue. Background, yellow. ACID FUCHSIN - PICROINDIGO CARMINE A cytological stain for root tips Solutions required: A. Acid Fuchsin 1% aqueous B. Saturated Picric acid, aqueous . . i volume Saturated Indigo carmine, aqueous i volume C. Acetic acid 0-25% aqueous Technique: 1. Bring sections through to distilled water in the usual manner. 2. Stain from five to twenty minutes in the acid fuchsin solution. 3. Rinse in water until stain ceases to come out of the sections. 4. Stain from five to fifteen minutes in solution B. 5. Wash with solution C. 6. Wash rapidly in 70% alcohol, in which the sections will appear red. 7. Wash rapidly in 95% alcohol, until sections appear greenish* 8. Dehydrate rapidly with two changes of absolute alcohol. 9. Clear in xylol and mount. Results: With onion root tips as an example, chromosomes and late prophase stages are in various shades of bright or dark red. Early prophases, bluish red. Nucleoli, clear blue. Spindle fibres and cell walls are stained a dark blue against a light cytoplasm. From Plant Microtechnique by D. A. Johansen, by courtesy of McCraw-Hill Book Company, Inc., New York. 290 SECTION FOUR ALIZARIN RED, S For mitochondria, cell inclusions, etc. (Benda's method) Solutions required: A. Chromic acid i% 30 ml. Osmic acid 2% . . 8 ml. Glacial acetic acid 6 drops B. Pyroligneous acid 100 ml. Chromic acid i % 100 ml. C. Potass, dichromate 2%. D. Iron Alum 4% aqueous. E. Alizarin Red S, saturated in abso- lute alcohol • • I ml. Distilled water • • 90 ml. F. Benda's crystal violet: Crystal violet, saturated in 70% alcohol . . • • 100 ml. Alcohol 70% • • 99 ml. Hydrochloric acid • • I ml. Aniline water • • 200 ml. Technique: 1. Fix for eight days in Solution A. 2. Wash for one hour in water; then transfer to Solution B for twenty-four hours. 3. Transfer to Solution C for twenty-four hours. 4. Wash for twenty-four hours in water. 5. Dehydrate; clear; embed in paraffin wax. 6. Mordant sections on slides for twenty-four hours with Solu- tion D ; then rinse in water. 7. Stain for twenty-four hours in Solution E. 8. Rinse with water; flood slides with Solution F and warm gently until vapour is given off; or stain at room temperature for twenty-four hours. 291 ( MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 9. Rinse in water; then differentiate for two to three minutes in 30% acetic acid until the nuclei appear reddish. 10. Wash in running water for five to ten minutes ; blot dry. 11. Dip into absolute alcohol; then pass through bergamot oil into xylol and mount. Results: Mitochondria, violet. Chromatin and archoplasm, brownish red. Certain secretion granules, pale violet. Centrosomes, reddish violet. ANILINE FUCHSIN - IODINE GREEN For mitochondria Solutions required: A. Acid fuchsin aqueous 10% . . 90 ml. Aniline oil. . . . . . . . 10 ml. Mix well by shaking at intervals over a period of several hours ; then filter. Note: This solution deteriorates after three or four weeks. B. Potassium permanganate 1% aqueous. C. Oxalic acid 5% aqueous. D. Iodine green 1% aqueous. Technique: 1. Small pieces of tissue are fixed in Regaud's Fluid for four days. 2. Transfer to 3% aqueous potassium dichromate for eight days changing the solution at intervals of two days. 3. Wash in running water overnight. 4. Dehydrate in alcohol as usual and clear in xylol. 5. Embed in paraffin wax; cut sections not more than 5/^ in thickness. 6. Fix sections to slides; pass through xylol and descending 292 SECTION FOUR grades of alcohol down to distilled water in the usual manner; then blot carefully. 7. Place the slides, sections facing upwards, on the corner of a tripod; flood the preparation with aniline fuchsin (Solution A) and heat gently with a small bunsen flame until vapour rises. 8. Remove the flame and allow the stain to act for five to ten minutes. 9. Pour oflF the excess stain and drain for a few seconds. 10. Rinse well in distilled water. 11. Immerse in the potassium permanganate for five seconds; then pour off^ excess. 12. Immerse in the oxalic acid solution for five seconds; then pour off excess. 13. Wash with distilled water; then stain with the iodine green solution for five to ten seconds. 14. Pour off excess stain; drain for a few seconds; dehydrate quickly with 95% and absolute alcohol. 15. Clear in xylol; mount in Cristalite. Results: • Mitochondria, crimson. Nuclei, green. ANILINE FUCHSIN - PICRIC ACID (Altmann) For mitochondria Solutions required: A. AltmanrCs Fluid: Potassium dichromate 5% aqueous i volume Osmic acid 2% aqueous . . i volume Note: Although the^ penetration power of this fi[xative is poor, it is very satisfactory for surface fixation. B. Acid fuchsin . . . . . . 20 gm. Aniline water . . . . • • 95 nal- C. Picric acid saturated in absolute alcohol. 293 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1 . Small pieces of tissue, not more than z mm. in diameter, are fixed for twenty-four hours in Altmann's Fluid. 2. Wash for an hour in running water ; then dehydrate ; clear, and embed in paraffin wax in the usual manner. 3. Sections, not thicker than 4//, are brought down to distilled water; then stained for six minutes in Solution B. 4. Pour off excess stain; then blot section carefully; then differentiate and counterstain by flooding the preparation with Solution C. 5. Rinse quickly in 95% alcohol; then dehydrate with absolute alcohol ; clear in xylol, and mount. Results: Mitochondria are stained crimson against a vivid yellow back- ground. 1 'j ANILINE SAFRANIN (Babe's) For demonstrating mitosis in animal tissues i Solutions required: < A. Aniline Safranin (Babe). s B. Alcohol 95% 99 ml. \ Acetic acid 1% . . . . . . i ml. j Technique: \ 1. Fix small pieces of tissue in Flemming's or Hermann's fluid and embed in paraffin wax. | 2. Stain sections for five to ten minutes in the safranin solution. 3. Rinse in water. 4. Differentiate in the acid alcohol (Solution B). 5. Rinse in 95% alcohol. 6. Rinse in absolute alcohol. 7. Clear in xylol and mount. Results: Mitotic figures are stained an intense red, while resting nuclei are deep pink to colourless. 294 SECTION FOUR BASIC FUCHSIN - PICRO INDIGOCARMINE (Alcoholic) For plant tissues, as a cytological stain for root tips Solutions required: A. Basic fuchsin i% in 70% alcohol. B. Indigocarmine . . . . . . 0-6 gm. Distilled water . . . . . • 50 ml. Picric acid, saturated aqueous 50 ml. Technique: 1. Sections are brought down to distilled water; then stained from ten to twenty minutes in the basic fuchsin solution. 2. Rinse in distilled water until the stain ceases to come out. 3. Stain from five to fifteen minutes in Solution B. 4. Rinse quickly in 70% alcohol until the sections appear red to the naked eye. 5. Rinse rapidly in 95% alcohol, followed by absolute alcohol until the preparation appears green to the naked eye. 6. Clear in xylol; then mount. Results: Chromosomes and late prophase stages, varying shades of red ; early prophases, bluish red; nucleoli, clear blue; spindle fibres and cell walls, dark blue; cytoplasm, light blue. BASIC FUCHSIN - PICRO INDIGOCARMINE (Aqueous) For chromosomes in plant tissues Solutions required: A. Basic fuchsin 1% aqueous. B. Picro indigocarmine. Technique: 1. Material is fixed in Nevashin or Regaud. 2. Wash in running water; dehydrate, clear and embed in paraffin wax in the usual manner. 3. Fix sections to slides and remove paraffin wax with xylol. X 295 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 4. Pass through absolute 90% and 70% alcohol down to distilled water, as usual. 5. Stain in the basic fuchsin solution for about five minutes. 6. Rinse in distilled water. 7. Stain in the picro indigocarmine solution for ten to twenty minutes. 8. Rinse in distilled water to which a few drops of hydrochloric acid have been added. 9. Differentiate in 80% alcohol for half to one minute, until red coloration ceases to come away. 10. Dehydrate with 70%, 90% and absolute alcohol. 11. Clear in xylol, and mount. Results: Chromosomes, brilliant red. Nucleoli and other cell com- ponents, sky blue. BREINL'S TRIPLE STAIN For chromosomes Solutions required: A. Iodine . . . . . . . . i gm. Potassium iodide . . . . . . 2 gm. Alcohol 90% . . . . . . 100 ml. B. Safranin 1% in 50% alcohol C. Methylene Blue, polychrome (Unna) D. Orange tannin Technique: 1. Fix sections to slides and remove paraffin wax with xylol as usual. 2. Rinse with two changes of absolute alcohol. 3. Mordant by immersing in solution A for fifteen minutes. 4. Rinse well with water. 5. Stain for at least half an hour in the safranin solution in a closed jar or tube. 296 SECTION FOUR 6. Wash well with water. 7. Stain with solution C for ten minutes. 8. Wash with water; drain and remove excess water, but do not allow the preparation to dry. 9. Place the slide under the microscope; then cover the pre- paration with orange tannin, by means of a pipette. 10. Observe the progress of the staining under the microscope until the orange tannin has replaced the blue in the cytoplasm. 1 1 . Withdraw excess fluid from the slide by means of a piece of filter paper to avoid the risk of spilling over the microscope stage and condenser. 12. Take the slide away from the microscope and wash well with 95% alcohol. 13. Dehydrate in absolute alcohol. 14. Clear and complete differentiation in aniline oil. 15. Rinse with cedarwood oil. 16. Drain and remove excess cedarwood oil from around the preparation by blotting or wiping with a clean fluffless duster. 17. Mount in Cristalite or Canada balsam in Xylol or Emexel mountant. Results: Chromosome threads, blue. Metaphase chromosomes, red. Cytoplasm, pale yellow. Reference: Bolles-Lee, nth edition, p. 664. COPPER CHROME HAEMATOXYLIN (Bensley) For mitochondria Solutions required: A. Altmann's Fluid B. Copper acetate, saturated aqueous C. Haematoxylin 10% in absolute alcohol ripened . . . . . . • . 5 ml. Distilled water . . . . . . 90 ml. 297 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES D. Potassium chromate neutral, 5% aqueous E. Weigerfs Borax Ferricyanide Mixture Borax 1% aqueous . . . . . . 100 ml. Potass, ferricyanide . . . . 1-25 gm. Technique: 1 . Fix very small pieces of tissue in Altmann's fluid from 1 2 to 24 hours. 2. Wash, dehydrate, clear and embed in paraffin wax in the usual way. 3. Cut sections from 4 to 5jLt in thickness and affix them to slides. 4. Remove paraffin wax with xylol. 5. Wash well with absolute alcohol. 6. Wash with 90% alcohol. 7. Wash with 70% alcohol. 8. Wash with distilled water. 9. Immerse in solution B for 5 minutes. 10. Wash with several changes of distilled water for a total time of one minute. 1 1 . Stain with the haematoxylin (solution C) for i minute. 12. Wash in distilled water. 13. Immerse in solution D for one minute, which should turn the sections dark blue-black : if they are only light blue in colour, rinse in distilled water and return to the copper acetate solution and if necessary repeat several times until the sections are dark blue-black after a minute in solution D, or until no increase in colour is obtained. 14. Rinse with distilled water. 15. Immerse in solution B again for a few seconds; then again rinse in distilled water. 16. Differentiate under the microscope with a mixture consist- ing of one volume of solution E and two volumes of distilled water. 17. Wash with tap water for 6-8 hours. 18. Dehydrate, clear and mount. 298 SECTION FOUR Results: Mitochondria are stained deep blue against a yellowish back- ground. COTTON RED - METHYL VIOLET - ORANGE G A cytological stain for plant tissues Solutions required: A. Cotton red i% aqueous. B. Methyl violet loB i% aqueous. C. Orange G - Clove oil. Technique: 1. Stain sections of Flemming-fixed material in the cotton red solution from eight to twenty-four hours. 2. Wash v/ell in water ; then stain in the methyl-violet solution from ten minutes to an hour according to the material. 3. Pour off excess stain; then wash in water. 4. Wash well with 95% alcohol; followed by thorough rinsing with absolute alcohol. 5. Drain well, and blot carefully. 6. Immerse in orange G - clove oil from five to ten seconds. 7. Pour off excess stain ; then wash the preparation with clove oil. 8. Differentiate in a fresh lot of clove oil for about one half to one minute, controlling by examination under the microscope until the violet stain is satisfactory. 9. Pour off excess clove oil; then wash with three changes of xylol. 10. Mount in Canada balsam in xylol. Results: Metaphase and anaphase chromatin, red. Prophase chromatin, violet. Chromoneata, red. Chromosome matrix, purple. Spindle fibres and plastids, violet. Cytoplasm, light brownish grey. Nucleoli, red. 299 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES FEULGEN STAIN For mitosis in plant cells Solutions required: A. Feulgen's fuchsin. B. Potass, metabisulphite io% . . 5 ml. N/i HCl 5 ml. Distilled water . . . . . . 100 ml. C. Fast green FCF 0-5% in 70% alcohol. Technique: Tissues should be fixed in Flemming and embedded in paraffin wax. 1. Sections are brought down to distilled water. 2. Rinse in N/i HCl; then transfer to N/i HCl at 60° C. for four to five minutes; afterwards rinsing in cold N/i HCl. 3. Rinse in distilled water; then stain for three to four hours in Solution A. 4. Drain and immediately transfer to a stoppered jar containing Solution B for ten minutes; then transfer to a second jar of Solu- tion B for ten minutes, followed by ten minutes in a third jar. Note: The jars must be kept closed. 5. Rinse in distilled water; then counterstain for one half to one minute in Solution C. 6. Dehydrate; clear and mount. Results: Chromosomes, reddish violet. GENTIAN VIOLET - PICRIC ACID - IODINE For chromosomes in plant cells Solutions required: A. Potassium iodide • • • • 0-5 gm Iodine, resublimed . . • • • • 0-5 gm Water • • . . 5 ml. Absolute alcohol . . . . , 45 ml. 300 SECTION FOUR Dissolve the potassium iodide in the water, then add the iodine and shake till dissolved, afterwards adding the alcohol and mixing well. B. Crystal violet i% aqueous. C. Picric acid 0*5% in absolute alcohol. Technique : 1. Fix in Navashin or Fleming. 2. Take the preparation (smear or section) through to 70% alcohol in the usual way. 3. Pass through 95% alcohol. 4. Mordant for fifteen minutes in solution A. 5. Rinse well in water. 6. Stain with solution B for ten to fifteen minutes. 7. Rinse well in water. 8. Immerse again in solution A for a few minutes. 9. Rinse in 95% alcohol. 10. Immerse the preparation for about one second in the picric acid solution. 11. Wash immediately with absolute alcohol for a few seconds. 12. Rinse in clove oil until the violet stain ceases to come out of the preparation. 13. Wash well in two changes of xylol. 14. Immerse in a jar of xylol for about an hour. 15. Drain off excess xylol and mount in Canada balsam in xylol or in Cristalite. Results: Chromosomes: rich purple, each chromosome being sharply defined, Cytoplasm: yellow. Note: This technique is an improvement on Newton's Gentian Violet - Iodine in that fading does not occur and sharp differentia- tion is obtained of Chromosomes that are close together. Reference: Smith, F. H. (1934), Stain Tech., 9 95-6. 301 SECTION FOUR HAEMATOXYLIN (Regaud) For mitochondria Solutions required: A. Formalin (40% formaldehyde) . . 25 ml. Potassium dichromate . . • • 3 gm- Distilled water . . . . . . 98 ml. B. Potassium dichromate 25% aqueous. C. Ammonia - ferric alum 10% aqueous. D. Haematoxylin (Regaud). E. Iron Alum 5% aqueous. Technique: 1. Small pieces of tissue are fixed for three to five days in Solu- tion A, which should be freshly prepared and changed each day. 2. Mordant by immersing for ten to fourteen days in Solution B. 3. Wash in running water for twenty-four hours. 4. Dehydrate by immersing in ascending grade of alcohol, and clear in the usual manner. 5. Embed in paraffin wax and cut sections no thicker than 5//. 6. Fix sections to slides ; de-wax and pass through descending grades of alcohol down to water. 7. Mordant sections for one to three days in a stoppered jar in the incubator at 37° C. 8. Rinse for five minutes or so in running water. 9. Immerse for twenty-four hours in the haematoxylin (Solu- tion D). 10. Differentiate in Solution E, controlling at intervals by examination under the microscope. 11. Rinse in running tap water for twenty to thirty minutes. 12. Drain off excess water; then rinse with 95% alcohol. 13. Dehydrate with absolute alcohol. 14. Clear in xylol and mount. 302 SECTION FOUR Results: Mitochondria are conspicuously stained intense black. HAEMATOXYLIN - SAFRANIN For differentiating nucleoli and other nuclear constituents in plant cells Solutions required: A. Iron Alum 3% B. Haematoxylin 10% in absolute alcohol, well ripened . . . . . . 5 ml. Distilled water . , . . • • 45 iiil* C. Safranin saturated in absolute alcohol . . . . . . . . I volume Aniline water . . . . . . i volume D. Picric acid 1% in 95% alcohol. Technique: 1. Fix sections to slide and take down to distilled water. 2. Mordant with solution A for two to three hours. 3. Wash in running water for five minutes. 4. Rinse in distilled water. 5. Immerse in the haematoxylin for an equal length of time. 6. Differentiate very carefully with solution A, controlling by examination under the microscope while the preparations are still wet. 7. When the nucleoli have been almost completely decolorized remove the Iron alum solution quickly by rinsing in water. 8. Wash in running water for an hour or longer. 9. Immerse in solution D for twelve to sixteen hours. 10. Differentiate for a few seconds in Picro-alcohol (solution D.) 1 1 . Wash with two changes of absolute alcohol. 303 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 12. Clear in xylol. 13. Mount in Cristalite or Canada balsam in xylol and examine under the oil immersion objective. Results: Nucleoli, brilliant red. Chromosomes during metaphase and anaphase are stained a bright red and stand out sharply against a dark background ; during prophase and telophase the chromosomes are considerably darker. Trabants are usually red and are attached to the chromosomes by a black thread. Note: This technique gives the best results after a killing fluid containing picric acid, but chromic acid fixatives also give good results. Reference : Plant Microtechnique (ist ed., p. 75) by D. A. Johansen, by courtesy of the McGraw-Hill Book Co. IRON ACETO CARMINE (Belling) For chromosomes in microsporocytes Solutions required: Iron Aceto Carmine [Belling) Acetic acid 50% aqueous . . . . 100 ml. Carmine, powdered . . . . i gm. Boil under reflux condenser for half an hour. Cool and filter. Add a few drops of ferric hydrate in 50% acetic acid. Technique: Temporary preparations : 1 . Anther smears are made on slides by teasing and squashing. 2. Place a drop of the stain on a smear ; then cover with a cover- slip. 3. Place slide, coverslip facing upwards over a corner of a tripod and heat gently until steam rises from the edges of the coverslip. 4. Examine under the microscope at once. 304 SECTION FOUR Results: Chromatin is stained deep translucent red, while cytoplasm is unstained. Permanent preparations : 5. Proceeding from Stage 4 (above), immerse the preparation in a Petri dish containing 10% acetic acid until the coverslip becomes loose. 6. Immerse the slide and the coverslip in a mixture consisting of equal volumes of absolute alcohol and glacial acetic acid, con- tained in another Petri dish. 7. Transfer to a dish containing a mixture consisting of three volumes of absolute alcohol and one volume of glacial acetic acid. 8. Transfer to a mixture consisting of one volume of glacial acetic acid and nine volumes of absolute alcohol. 9. Transfer to absolute alcohol ; then into two changes of xylol. 10. Refix the coverslip to the slide with a drop of Cristalite. METHYL GREEN - ACID FUCHSIN - ERYTHROSIN A cytological stain for plant cells Solutions required: A. Methyl Green 1% aqueous. B. Acid fuchsin 1% aqueous. C. Erythrosin 1% aqueous. Technique: 1. Take sections down to distilled water in the usual manner. 2. Stain for an hour in solution A. 3. Remove excess stain by washing thoroughly with water. 4. Stain in the acid fuchsin solution for one minute. 5. Wash well with distilled water to remove excess stain. 6. Stain with the er)rthrosin solution for two or three seconds. 7. Wash well with distilled water and drain and blot off excess water, but do not allow the section to dry. 305 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 8. Rinse with 70% alcohol. 9. Rinse quickly with 90% alcohol. 10. Dehydrate quickly but thoroughly with two changes of absolute alcohol. 11. Clear in xylol and mount in Canada balsam in xylol or in Emexel. Results: Chromatin granules and nucleoli in early stages of micro- sporogenesis are stained green while the linin threads are red. Chromosomes are stained brilliant green in later stages. Reference: Cooper, D. C. (1931) American J. Bot., i8, 337. METHYL GREEN - ACID FUCHSIN For chromosomes, etc., in plant tissue Solutions required: A. Methyl green 2% aqueous . . 100 ml. Acetic acid 1% . . . . . . 2 ml. B. Acid fuchsin 1% aqueous. Technique: 1. Sections are brought down to water; then stained from six to seven hours in the methyl green solution. 2. Wash in water until the stain is almost entirely removed from the non-lignified elements (this process should be controlled by examination under the microscope v/hile the preparation is still wet). 3. Rinse in water. 4. Stain from five to ten minutes in the acid fuchsin solution, controlling under the microscope to ensure that the green is not extracted from the lignified tissues. 5. Wash rapidly in 95% alcohol, followed by absolute alcohol. 6. Clear in clove oil ; then pass through xylol and mount. Results: Cytoplasm and plastin, light red; chromosomes and nuclei, green. 306 SECTION FOUR METHYL VIOLET - EOSIN SCARLET A botanical stain for mitosis in root tips Solutions required: A. Methyl Violet 5B aqueous 1%. B. Picric acid 1% in absolute alcohol. C. Eosin scarlet, saturated in equal volumes of clove oil and absolute alcohol. Technique: 1. Sections are fixed to slides and taken down to water in the usual manner. 2. Stain for twenty to thirty minutes in the methyl violet solu- tion ; then wash with distilled water. 3. Differentiate with the picric acid solution for a few seconds. 4. Immerse in absolute alcohol to which has been added 0-15% strong ammonia solution, for fifteen to twenty seconds. 5. Immerse in absolute alcohol for ten to twenty seconds. 6. Counterstain for five to fifteen seconds in the eosin scarlet solution. 7. Clear in clove oil ; rinse with xylol ; mount. Results: Resting and dividing chromatin, purple. Cytoplasm, pink. Cell walls, red. Plastin, deep red. NIGROSINE For the study of salivary chromosomes of Drosphila Solutions required: A. Acetic acid 45% aqueous B. Absolute alcohol . . . . . . 70 ml. Distilled water . . . . . . 29 ml. Hydrochloric acid, pure, cone. . . i ml. 307 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES C. Nigrosine, alcoholic (Michrome brand) . . . . . . . . i gm. 70% alcohol 100 ml. Technique : 1. Dissect the specimen in 45% acetic acid. 2. Treat with solution B for one to two minutes, keeping glands well covered with the reagent. 3. Wash the specimen carefully two or three times with distilled water, after which a large drop of water is left on the specimen for five minutes. 4. Drain thoroughly and carefully. 5. Add a drop of the nigrosine solution. 6. Place a coverslip over the preparation but do not squash. 7. Allow the stain to act for ten to fifteen minutes. 8. Place a piece of blotting paper over the coverslip and slide and squash the preparation. 9. Immediately place a large drop of glycerine on one edge of the coverslip. 10. The preparation may now be examined at once, but after about twenty-four hours the glycerine will have penetrated the specimen thoroughly, when it is recommended, for the sake of convenience, to remove the excess liquid and ring the coverslip with wax. Note: Preparations obtained as above will last for reasonably long periods provided they are carefully handled. Results: Salivary glands specifically and intensely stained ; only the outer bands being coloured, except in heterochromatin regions. When the preparation is ageing, however, the interbands as well as the cytoplasm, become faintly stained. Reference: Pares, Ramon (1953), Nature, 172, no. 4390. 308 SECTION FOUR SAFRANIN CRYSTAL VIOLET (Hermann) For chromosomes Solution required: A. Safranin . . . . . . . . i gm. Aniline water . . . . • . 50 ml. Absolute alcohol . . . . • • 50 ml. B. Crystal violet 1% in 70% alcohol. Technique: 1. Stain for three to twenty-four hours in the safranin solution in a closed jar or tube. 2. Destain briefly with 50% alcohol. 3. Counterstain with the crystal violet from thirty seconds to five minutes. 4. Rinse quickly with 90% followed by absolute alcohol. 5. Clear with clove oil. 6. Wash with xylol, and mount. Results: Chromosomes and nucleoli are stained red; resting nuclei and chromatin granules violet: propase and early telophase nuclei may show violet chromonema and red chromatin granules ; cyto- plasm, light violet; spindle, deeper violet. Reference: Hermann, Arch, Mik. Anat., 34, 58. SAFRANIN - GENTIAN VIOLET - ORANGE G. (Flemming Tricolour stain) For chromosomes, etc. Solutions required: A. Safranin 3*5% in 95% alcohol . . 10 ml. Aniline water . . . . . . 10 ml. B. Gentian violet 5% aqueous. C. Orange G 1% aqueous. 309 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique: 1. Sections of material, which have previously been fixed in Flemming's fixative, are stained for two to twenty-four hours with the safranin solution, in a stoppered staining jar. 2. Differentiate in acid alcohol until colour almost ceases to come out of the sections and microscopic examination shows the chromosomes in metaphase stained clearly with the safranin. 3. Rinse with distilled water. 4. Immerse in the gentian violet solution, in a staining jar for one half to three hours, washing with water at intervals and examining under the microscope until the prophase chromatin is clearly stained violet. 5. Differentiate in the orange G solution for thirty to sixty seconds when pale violet clouds should be given off. 6. Rinse in absolute alcohol until scarcely any more colour comes out of the sections. 7. Differentiate in clove oil and while very faint clouds of colour are still coming away, rinse with benzol and mount in balsam. Results: Chromosomes, red. Nucleoli, red. Metachromatic chromatin, deep purple. Cytoplasm, orange. Spindle, blue-violet. Note: The action of this stain should be so precise that in one strand of chromatin the diffused portion will take the violet, while the condensed section should be stained with the safranin. WRIGHT'S STAIN For chloroplasts in tissue spreads and for plant cytology Note: Valuable for showing cytoplasmic changes in various tissues and cell inclusions. Reveals cytoplasmic differentiations and experi- mental change. The method is generally applicable for cytological work wherever material may be spread (not smeared) and dried rapidly. Produces excellent preparations of chloroplasts. Where cells separate well and where there is neither an excess of water in tissue fluid nor concentrated protein scum, conditions are favourable for a good spread of uninjured cells and an excellent stain. Solution required: Wright's stain. 310 SECTION FOUR Technique: 1. A Nitella cell, carefully isolated from the plant, is placed on a piece of filter paper from which it is transferred to a clean dry slide. 2. The uninjured cell is pricked so that it bursts out its sap and the chloroplasts in effective spread. 3. Lift the deflated cell with needles and transfer to an adjacent dry region and tear to release more chloroplasts with a small amount of sap. Arrange areas of the cell wall smoothly so that no portion of the cell or its contents are lost in the preparation. By spreading the cell fluid about in this way there is no excess in any region. 4. Dry rapidly with gentle heat. 5. Place I ml. of Wright's stain on the dried spread, and leave it to act for one minute ; then add 2 ml. distilled water and rock the slide gently to ensure thorough mixing. 6. Allow this diluted stain to act for three to five minutes ; then pour off and wash with distilled water until the thin portion of the films appears pink to the naked eye ; then pour off excess stain. 7. Wash with distilled water and allow the preparation to dry before examining. Results: Chloroplasts of normal Nitella are basophilic and of marked alveolar structure. Large, homogeneous-appearing vacuole plas- tids which are noted in the streaming of the living cell vacuole are brilliantly eosinophilic. However, they prove to be complex with a central raphe, usually colourless arrangements of both basophilic and eosinophilic granules; and both bilaterally distri- buted lacunae varying according to experimental conditions from colourless to brilliant blue. The staining of Tunicate material is so critical that species differences are readily noted in comparative studies of the same cells. Reference: Koehring, Vera (1951), Stain Tech., 26, 29. 311 SECTION 5- FLUORESCENCE MICROSCOPY (a) GENERAL INFORMATION Fluorescence is the property possessed by many substances of converting short wavelengths of Hght into longer wavelengths. In microscopy the substances and structures of most interest are those which convert ultraviolet light into light of the visible spec- trum, as only substances of this character can be observed directly. It is, of course, well known that most living organisms are pro- foundly affected by short light waves, and a great deal of infor- mation as to their structure has been obtained by the study of the appearance of these organisms under the influence of invisible light rays. If individual cells or structural units are examined before, during, and after ultraviolet treatment, enough of this effect should be discovered to impart some under- standing as to the changes which occur in the animal, or plant, as a whole. In fluorescence microscopy structural details are rendered visible by: {a) innate auto-fluorescence, a property possessed by most tissues which when excited by short light waves become clearly visible since they become luminous and glow or " fluor- esce " with a radiance of their own, or by {h) secondary fluorescence which is known as " fluorochromy " and is brought about by the process of treatment of the tissues with fluorescent dyes and cer- tain alkaloids (e.g. Berberine sulphate) and other substances. It is proposed to deal only with fluorochromy in the short space of this chapter. Fluorescent dyes and other substances used for this purpose are known collectively as " fluorochromes " ; these materials are selectively absorbed by certain parts of the cell. When tissues, bacteria or protozoa, which have undergone treatment with fluoro- chromes are examined under the microscope, using ultraviolet light instead of transmitted light of the visible spectrum, they become visible as bright luminous objects against a dark back- ground. Cells stained with fluorochromes absorb the ultraviolet rays of short wavelength, and emit this energy in the form of 315 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES fluorescent light in the visible spectrum. Basic and acidic fluoro- chromes act specifically to stain certain cellular structures as do the more common microscopic stains such as, for instance, methylene blue and eosin. The colour and the intensity of fluorescence depends on the relative basophilia and acidophilia of the individual cells, and upon the nature of the particular fluoro- chrome. Fluorochromy may be employed with advantage in the study of living organisms: for instance, uranin, a non-toxic stain, may be injected into mice and frogs and the living organs can be studied without interfering with their functioning. Fluorochromy is also of practical importance for the demonstration of diphtheria bacilli, tubercle and leprobacilli, malaria, etc., as well as for virus research. (b) EQUIPMENT REQUIRED Contrary to the generally held belief, the apparatus required for fluorescence microscopy is fairly simple and inexpensive. A special microscope is not required. 1. B.T.H. Mazda Mercury Vapour Lamp (box type) ME 250 w/50/5. 2. A simple convex lens to project the image of the light source through a suitable blue filter to the microscope mirror. The lens and light source should be encased with a black hood to prevent scattering of the rays. 3. A yellow filter which is placed in the oculars of the micro- scope to prevent any harmful effect of ultraviolet light to the microscopist's eyes. For this purpose Ilford's minus blue Micro 4 is recommended. It is, of course, essential that fluorescence microscopical examin- ations must be carried out in a darkroom to be successful. It has been stated that microscope slides of special glass are necessary for fluorescence microscopy, but provided they are not more than 1-3 mm. in thickness ordinary microscopic slides have been found quite satisfactory. 316 SECTION FIVE (c) STAINING METHODS Notes on fluorochromic staining technique: (a) Fixatives containing salts of heavy metals (with the exception of zinc) ; chlorine, bromine, iodine, and nitro compounds should be avoided as these exert a quenching effect on fluorescence. The most suitable fixatives are 5 to 10% formalin, or Kahle's fluid. (b) If tissues are embedded in paraffin wax, all traces of the wax, which is auto-fluorescent, must be removed before sections are stained and examined. (c) A special grade of immersion oil known as fluoroil, which is non-fluorescent, should be used for high-power examination, since cedarwood oil and most of the immersion oils available for ordinary microscopy are unsuitable for fluorescence work. (d) The usual mounting media, as used for ordinary micro- scopy, contain highly fluorescent materials which render them un- suitable for fluorescence work, and should, therefore, be avoided. For temporary mounts, glycerine may be used, and for permanent mounts there is a satisfactory medium on the market under the name of Fluormount. Fluorochromes, of which auramine O, coriphosphine, acridine yellow Hi 07, aesculine, acridine orange, primuline, thiazole yellow are examples, are frequently used in very dilute solutions to pro- duce characteristic fluorescent colours, and when preparations which have been treated with these solutions are examined in trans- mitted light of the visible spectrum, they appear to be unstained or only very faintly tinted. Some explanation as to the colour differentiation obtained by the use of general-purpose fluorochrome of which acridine yellow Hi 07 is an example, is explained by the fact that fluorescence colour is effected by hydrogen-ion concentration, and as fluoro- chromes also exhibit differential absorption by various tissues, the production of a great variety of fluorescent colours is brought about by the influence of these two factors. Nuclei can be differ- entiated from cytoplasm by the use of any one of the following general fluorochromes: acridine yellow Hi 07, coriphosphine, phosphine 3R, or acriflavine. Titan yellow, rhodamine B, phosphine 3R, methylene blue are 317 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES all excellent fluorochromes for fat, while berberine sulphate is much used for protozoal parasites, particularly for malaria. Thio- flavine has been found satisfactory for virus and for bacteria, as well as for general fluorochromic staining, while acriflavine has been used for trypanosomes and as a general stain, and uranin is one of the most suitable fluorochromes for intravital staining. All these fluorochromes are used in very dilute aqueous solutions, that is to say, something to the order of o-i to o-oi per cent. I. Differential Staining of C3rtoplasni, Nuclei, Nucleoli and Chromatin Solutions required: A. Congo red o-i% aqueous or acid fuchsin 0'i% aqueous. B. Acridine yellow Hi 07 o-i% aqueous or coriphosphine o-i% aqueous. Technique: 1. Sections are stained for two minutes with the Congo red or acid fuchsin solution. 2. Pour off excess Congo red ; then stain for two minutes with the acridine yellow or coriphosphine solution. This technique gives a very sharp differentiation. 2. Staining of Fat Method A. Solution required: Phosphine 3R o-i% aqueous, or methylene blue 1% aqueous. Technique: 1. Stain frozen section for two minutes ; then rinse in water. 2. Examine in water. Results: With phosphine 3R fat is observed as a silver fluorescence 318 SECTION FIVE against a brown background and due to the omission of lipid sol- vents in this technique, smaller and more numerous fat droplets can be seen than in the case of the sudan techniques as used in ordinary microscopy. With methylene blue fat gives a blue fluorescence. Method B. Solution required: Thioflavine S i% aqueous. Technique: 1. Frozen sections are stained for two minutes; then rinsed in water. 2. Examine in water. Results: Fats appear violet against a dark background. Note: With this method fewer lipids are stained than with phosphine 3R. 3. Intravital Staining with Fluorochromes Dyes used for this purpose must be water soluble, non-diffus- ible in the living body, non-toxic in the workable dilutions re- quired, and highly fluorescent even in greatly diluted solutions. Uranin possesses all these qualifications and is one of the most useful fluorescent dyes for intravital work, as stated earlier in this chapter. Acriflavine is another useful dye for this purpose, al- though it is not so intensely fluorescent as uranin. The fluor- escence of uranin is impaired in basic solution so that it appears most readily in organs of an acid reaction. It is used in o-i% solution in physiological saline, in which form it should be injected into the animal's blood stream or into the organ to be studied. The colour of its fluorescence varies with hydrogen-ion changes and consequently it is of great value as an intravital hydrogen-ion indicator. The colour changes are easily visible in dilutions to the order of one part in ten millions and in dilute solutions the inten- sity of the fluorescence has a definite relation to the concentration of the dye and consequently the intensity of the fluorescence 319 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES serves as an indicator of the amount of uranin present. Primulin, thioflavine, rhodamine B, berberine sulphate have also been found useful for intravital work. Many important discoveries in the biological field are due to fluorescence microscopy and although a great many beautiful and highly contrasting colour combinations have been obtained by the techniques devised up to the present time, fluorescence microscopy has scarcely yet emerged from the embryonic stage, and there is tremendous scope for experiment and improvement both in regard to technique and apparatus. 4. Staining of Muscle Solutions required: A. Primulin o-i% aqueous. B. Sodium nitrite 2% aqueous. C. Hydrochloric acid, cone. . . 10 ml. Distilled water . . . . . . 90 ml. D. Resorcinol, saturated aqueous solution. Technique: 1. Sections are stained for two to five minutes with primulin solution. 2. Wash quickly; then immediately transfer to the sodium nitrite to which has previously been added, with stirring, an equal volume of the diluted hydrochloric acid (Solution C above), and leave therein for fifteen to twenty minutes. 3. Wash in water; then immerse for fifteen to thirty seconds in the resorcinol solution. Results: Muscle tissue fluoresces with a strikingly luminous green fluorescence. 320 SECTION FIVE 5. Differentiation of Nerve Tissues Method A. Solution required: Acridine yellow Hi 07 aqueous o- 1%. Technique: 1. Sections are stained with the acridine yellow for two minutes. 2. Pour off excess stain and rinse with distilled water. 3. Dehydrate in the usual manner; clear in xylol; mount in Fluormount. Results: Neurophile nerve tissue appears light blue, while cortical nerve tissue is light yellow to orange. Method B. Solution required: Acridine orange o-i% aqueous. Technique: 1. Sections are stained with the acridine orange solution for two minutes. 2. Pour off excess stain, rinse with distilled water. 3. Dehydrate as usual, clear in xylol in Fluormount. Results: Neurophile is bluish-grey while cortical tissue is brownish orange. Method C. Solution required: Acriflavine o-i% aqueous. Technique: 1. Stain with the acriflavine solution for two minutes; then pour off excess and rinse in water. 2. Dehydrate rapidly ; clear in xylol and mount in Fluormount. Results: Neurophile is blue, while cortical tissue appears brownish yellow. 321 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 6. Staining and counting of bacteria, and differentiating living from dead cells, in soil Solutions required: A. Buffer solution pH 7 B. Solution A . . Acridine Orange, FS C. Solution A Solution B D. Solution A Solution B E. Solution A Solution B F. Solution A Solution B G. Solution A Solution B 200 ml. 100 ml. o-i gm. 5 ml. 5 ml. 5 ml. 10 ml. 3 ml. 9 ml. 2 ml. 8 ml. I ml. 9 ml. Technique : 1. Place I gm. of the sifted soil in each of six test tubes, and label them consecutively B, C, D, E, F, and G. 2. Add 10 ml. of the appropriate solution to the five tubes labelled B, C, D, E, and F (i.e. 10 ml. solution B to tube B; 10 ml. solution C to tube C, etc.). Note: As soils differ in their ability to absorb the stain, the same soil must be stained with different concentrations of the acridine orange to ascertain the most suitable concentration. Solution G will be used for differentiating between living and dead bacteria. 3 . Shake each of the five tubes thoroughly ; then leave them to stand for five or ten minutes. 4. The most suitable concentration for the particular soil is that contained in the tube, which shows the least amount, compared with the others, of excess dye, and this specimen should be used for examination as follows: (a) For qualitative analysis of autochthonic bacteria : The dyed soil suspension is strongly centrifuged ; the upper (liquid) layer is poured off and discarded, leaving a small quantity 322 SECTION FIVE of sediment which is then well mixed with a drop of paraffin oil and covered with a coverslip before examination with the oil immersion lens. (a) For bacterial counts: A loopful of the stained and shaken soil suspension is placed on a slide, covered with a coverslip, and examined with an oil immersion lens. A counter of 20/x depth combined with a counting ruled eyepiece is used. The particles of soil covered with humus and the particles of humus slime show a dim red fluorescence and the living bacteria are green. The stained bacteria are not killed and may be used for culturing. To diff"erentiate between living and dead cells, solution G should be used: living cytoplasm and nuclei show green fluor- escence, whilst dead protoplasm develops a bright copper- coloured fluorescence. Reference: Strugger, S. (1948), Canad.J. Res. Sec. C, 26, pp. 188-93. 7. Vital Staining of Living Trypanosomes in Blood (S. Strugger 's Method) Solution required: Acridine orange o-oi% in 0-85% sodium chloride. Technique: 1. A drop of blood suspected of containing trypanosomes is mixed on a slide with a few drops of the acridine orange solution, and covered with a coverslip. 2. Examine with a blue light fluorescence microscope, which may be constructed as follows : parallel rays produced by attach- ing a convex lens to the lamp are filtered with a curvette (2 J cm. thick) filled with saturated solution of copper oxide ammonia so that only the blue light reaches the plane mirror of the microscope. A filter containing an orange glass which absorbs blue light quantitatively, but allows green, yellow and red to pass unchanged, is placed over the ocular. A slide with powdered anthracene in liquid paraffin is used for focusing. 323 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Results: Trypanosomes and leucocytes are seen as motile, bright, green luminous bodies while the erythrocytes are scarcely visible and are non-fluorescent. 8. Staining of Trypanosomes in Dried Blood Films Solution required: Auramine O o-i % aqueous . . lOO ml. Phenol crystals . . . . • • 5 gm. Technique: 1. Fix air-dried blood films, suspected of containing trypano- somes, in pure methyl alcohol for two to five minutes, then wash with distilled water. 2. Stain for about four minutes in the auramine solution then wash with distilled water for one to two minutes, keeping the slide in the dark. Results: Examined with blue light fluorescence microscope, erythrocytes appear as dark green, slightly luminous circles, while trypano- somes shine with a bright golden fluorescence against a black back- ground. By covering the film with a drop of liquid paraffin and a coverglass the contrast will be even more marked. 9. Staining of Tubercle Bacilli Solutions required: A. Auramine O aqueous 0-1% ., 100 ml. Phenol crystals . . . . • • 5 gm. Dissolve by shaking. Do not apply heat as aura- mine decomposes at 40° C. B. Methylene blue 0-1% aqueous. C. Potassium permanganate 0'i% aqueous. 324 SECTION FIVE Technique: Place sandalwood oil or fluoroil between the slide and condenser. Three sides of the microscope should be enclosed with a shield to exclude extraneous light. Sections: 1. Fix tissues in 5 to 10% formalin for twelve to twenty-four hours; embed in paraffin wax and cut sections 5 to lo// in thick- ness. 2. De-wax as usual, taking care that all traces of wax are com- pletely and finally removed from sections. 3. Bring sections down to distilled water as usual; then flood with the auramine solution and warm (not over 40° C.) for five to ten minutes, afterwards washing with distilled water. 4. Counterstain in methylene blue solution for thirty seconds; then rinse in distilled water, dehydrate as usual ; clear in xylol and mount in Fluormount. Smears: 1. Stain for five minutes in the auramine solution, afterwards washing with tap water and decolorizing in 25% sulphuric acid. 2. Immerse in the potassium permanganate solution for about thirty seconds to overcome any interfering fluorescence. Results: Tubercle bacilli appear as thin shining, slightly curved rods against a dark background. An Improved Method of Staining Tubercle Bacilli (H. Lempert's Method) Solutions required: A. Phenol 3% in distilled water . . 100 ml. Auramine O . . . . . . 0-3 gm. Dissolve by shaking. Do not apply heat, as aura- mine decomposes when heated above 40° C. 325 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES B. Hydrochloride acid, pure . . 0-5 ml. Sodium chloride . . . . 0-5 gm. Methyl alcohol, pure . . . . 75 ml. Distilled water . . . . • • 25 ml. C. Potassium permanganate o*i% aqueous. Technique: 1 . Smears of sputum or centrifuged urine deposits are fixed by heat in the usual manner. 2. Immerse the smears in the auramine solution for eight to ten minutes, afterwards washing them with tap water. 3. Decolorize by immersing for two minutes in each of two changes of Solution B; then wash in tap water. 4. Immerse in the potassium permanganate solution for about thirty seconds ; then wash with tap water ; blot and dry. Note: It is claimed that this method gives more positives than the Ziehl-Neelsen technique and there is a saving in time. Using Lempert's method, tubercle bacilli are visible under the two-thirds or the quarter-inch objective with a X 8 eyepiece, and the one- sixth-inch objective may be used for confirmation. 10. Staining Virus with Primulin Solution required: Primulin o-i% aqueous. Technique: 1. Stain for thirty seconds. 2. Mount if desired in a medium consisting of 100 gm. of best pale gum acacia dissolved in 100 ml. water and 50 ml. glycerine with the addition of 10 gm. chloral hydrate. Note: With primulin, which may also be used as a general fluorochrome, a blue-violet fluorescence is obtained. 326 SECTION FIVE ADDITIONAL REFERENCES Cowdrey, E. V. (1948), Lab. Tech. in Biol, and Med., 98, 2nd edition. Ellinger, P. (1940), " The site of acidification of urine in frog's and rat's kid- neys ", Quart, jf. Exp. Physiol.^ 30, 205-18. Ellinger, P. (1940), " Fluorescence microscopy in biology ", Biol. Rev., 15, 323-50. Evans, G. H., and Singer, E. (1941), " Fluorescence microscopy applied to ocular tissues ", Arch. Ophth., 25, 1007-19. Lempert, H. (1944), Lancet, 247, 818-22. Metcalf, R. L., and Patton, R. L. (1944), ** Fluorescence microscopy applied to entomology and allied fields," Stain Techn., 19, i. Strugger, S. (1948), " Fluorescence Microscope Examination of Trypano- somes in Blood." Canad. J. Res., Sec. E, 26, 229-31. SECTION 6- HISTOCHEMICAL METHODS MICRO - INCINERATION Micro-incineration is a technique employed to ascertain the relative positions occupied by inorganic salts in fixed tissues, and the total distribution of these mineral substances in cells and tissues. There are two methods of fixation, neither of which is perfect, namely; (a) Freeze- dry and (b) Chemical fixation. The first method requires reagents and apparatus which are not normally available in the average laboratory; also the method is exceedingly laborious and time consuming: there is a certain element of risk of injury to the operator who may not be accustomed to handling such substances as liquid nitrogen at a temperature of — i7o°C. Moreover, it is extremely doubtful if the elaborate *' Freeze-Dry " method offers any more accurate results than those obtained by the relatively simple chemical fixation method. By employing either method the total mineral distribution can- not be determined with an error of less than, say, io% as all minerals will not survive the incineration processes, and it is, therefore, proposed to describe here the more simple and straight- forward chemical method of fixation which can be carried out in any laboratory. Solution required: Absolute alcohol . . . . . . 90 ml. Formaldehyde 40% . . .. .. 10 ml. Note: It is best to use freshly distilled alcohol. On no account must the alcohol have been dried with copper sulphate nor any other mineral reagents. The formaldehyde must not have been treated with calcium carbonate, buffer salts or any other mineral reagents. It is essential that all the reagents used must be free from, and remain free from, dust and mineral matter; this refers also to the xylol and the paraffin wax. All aparatus used must be scrupulously clean, washed out with several lots of distilled water, partially dried with absolute alcohol, and in all cases, where at all possible, finally dried with a clean dust-free cloth. 331 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 1 . Fix small pieces of tissue in absolute alcohol, with a mech- anical agitator, for twelve to twenty-four hours, changing the alcohol at intervals of one hour during the day. 2. Clear in freshly filtered xylol. 3. Infiltrate with and embed in wax, which must not be plunged into water to hasten cooling. 4. Clean slides, in readiness to take the sections, by washing them several times in distilled water, then partially drying them with absolute alcohol, and finally drying them with a dust-free cloth. Store the slides away from dust and mineral matter until they are required for use. 5. Cut serial sections 3 to ^fi in thickness, taking particular care at this stage that they are not contaminated with dust or mineral matter. 6. Press serial sections onto slides, without any fixative or spreading agent, with the fingers which have been washed and dried with absolute alcohol. Note: It is now, at this stage, more than ever necessary to guard against contamination by dust. 7. Mount alternate serial sections on albuminized slides for normal histological staining and comparison later with the incinerated specimen. 8. The slides bearing the sections for incineration are placed in a quartz-tube electric furnace, but if that is not available a muffle furnace, the inside of which is entirely free from dust and debris, will serve the purpose. 9. Gradually raise the temperature so that it reaches a maximum of 60° C. at the end of the first three minutes. 10. During the next three minutes, gradually raise the tempera- ture to 70° C. 1 1 . Raise to 80° C. during the next two minutes. 12. Raise to 200° C. during the next five minutes. 13. During the next twenty-five minutes raise the temperature at the rate of 90° C. per five minutes, when the maximum of 650 ° C. will be attained. 332 SECTION SIX 14. Turn off the heat and allow the slides to cool in the oven for about thirty minutes. 15. Take the slides out of the furnace and lay coverslips over the sections, with a pair of forceps. 16. Seal the coverslips with De Kotchinsky's cement. 17. De-wax and stain the control serial sections, which have been set aside for the purpose, with suitable stains. 18. Examine the unstained, incinerated specimens comparing them with control serial sections. This can be most conveniently accomplished by employing two identical microscopes connected with a comparing ocular, the microscope with the normally stained sections being illuminated with ordinary bright-light condenser, and the one with the incinerated sections with a dark-field illumi- nator : this method facilitates the location of the same structure in both types of sections. Notes: There are many methods, some of which are given in this book {see index) of identifying the numerous minerals which might be present, but it is not possible to give them all here without entering the realms of Mineralogy, Fluorescence Analysis and kindred subjects which are foreign to a book of this kind, but which are well catered for in other text-books to which the reader is referred as well as to the various references given in McLung^s Handbook of Microscopical Technique^ Laboratory Technique in Biology and Medicine, by E. V. Cowdry, and the methods given in Microscopic Histochemistry by G. Gomori. 3.HYDROXY - 2-NAPHTHOIC ACID - TETRAZOTIZED o-ANISIDINE For demonstrating sites of carbonyl activity in frozen sections Solutions required: Important : All alcohol used in this technique must be aldehyde free. A. 3-hydroxy - 2-naphthionic acid hydrazide o-i gm. 50% alcohol . . 95 ml. Acetic acid glacial 5% . . . . • • 5 ml. 333 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES B. Absolute alcohol . . . . . . 30 ml. Phosphate buffer 1/15 M (pH 7-2). . 10 ml. Distilled water . . . . . . 20 ml. Tetrazotized o-anisidine . . . . o-o6 gm. Technique: 1. Tissues are fixed in 10% formalin (or 5% formol saline). 2. Without washing out fixative, cut frozen sections and place them on slides. 3. Allow the sections to become affixed to slides by drying in the atmosphere; then wash out formalin with several changes of water. 4. Immerse in solution B for one or more hours in an incubator at room temperature. 5. Remove excess reagent (solution B) by washing for two hours in 50% alcohol (aldehyde free). 6. Wash in several changes of water for several hours. 7. Immerse in solution C for one to two minutes. 8. Wash in several changes of distilled water to which a few drops of acetic acid have been added. 9. Drain off excess liquid; then mount in Glycerin Jelly. Results: Sites of carbonyl activity are indicated by the production of a blue pigment. Reference: Seligman, A. M. and Ashbel, R. (195 1), Cancer, 4, 579-96. METANIL YELLOW - IRON HAEMATOXYLIN For radioautographs prepared by mounting tissue sections directly onto photographic emulsions for study of thyroid carcinoma and all organs of man, as well as laboratory animals to which radio isotopes have been administered Solutions required: A. Metanil Yellow 0-25% aqueous. 334 SECTION SIX B. Haematoxylin . . . . . . i gm. Ethyl alcohol 95% . . . . . . 100 ml. C. Ferric chloride hydrated 50% aqueous . . . . . . • • 5 "nl* Distilled water . . . . . . 95 ml. D. Solution B . . . . . . . . i volume Solution C . . . . . . . . I volume Mix well, and allow to stand for two or three weeks before use. Filter immediately be- fore use. Note: The fresh solution must not he used as it darkens the emulsion gelatine ; 60 ml. of this solution is sufficient for about twenty radio- autographs. E. Acetone and Xylol, equal volumes Technique: 1. Surgical and post-mortem specimens from patients having recently received radioiodine (P^^) and tissues from laboratory animals to which radioiodine, radiosulphur (S^^) or radiophos- phorus (P22) has been administered, are fixed in Bouin's fluid. 2. Embed in paraffin wax, using the Dioxan technique. 3. Cut sections 7 to lo/x in thickness. 4. Transfer the section ribbons to the darkroom, place in a waterbath, and float them onto a photographic plate. (X-ray film, Kodak Medium lantern slides or Kodak nuclear track plates, are suitable for this purpose.) 5. Allow the sections to dry on the plate, when they will become permanently attached to the photographic emulsion. Note: For each isotype the processing fluids should be examined by a Geiger counter to ensure that there is no significant loss of the radioactive material from the tissue. 6. After proper photographic exposure, removal of the paraffin wax, development, fixation, washing and drying, the autograph is stained as follows: 7. Stain with metanil yellow (solution A) for five to fifteen seconds. 335 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 8. Wash in tap water until the slide is pale yellow: this should take about fifteen seconds. 9. Stain in solution D for one to three minutes. ID. Rinse with three changes of 70% alcohol for five seconds in each. 11. Wash for five seconds in 95% alcohol 12. Wash with acetone for five seconds. 13. Wash with solution E (acetone-xylol) ; for five seconds. 14. Wash with three changes of xylol. 15. Mount synthetic resins; e.g. D.P.X. or Cristalite. Results: Colloid, cytoplasm and connective tissue elements are stained from light yellow to light brown. Cell nuclei are stained deep- blue to black. Sites of radioactivity are indicated by blackening of the photo- graphic emulsion. Reference: Simmel, Eva B., Fitzgerald, P. J. and Godwin, J. T. (195 1), Stain Tech., 26, 25-8. METHYL - GREEN - PYRONIN - RIBONUCLEASE (Brachet) For detecting ribonucleic acid and desoxyribonucleic acid in the same cell Solutions required: I. Zenker's fixative containing 5% acetic acid or Carnoy Fixative UT Serra Fixative: Absolute alcohol Formalin (40% formaldehyde) Acetic Acid, glacial . . 60 ml 30 ml 10 ml 336 SECTION SIX 2.* Methyl green-Pyronin {Bracket) Methyl green, 00 chloroform washed 0-15 gm. Pyronin G . . . . . . . . 0-25 gm. Alcohol 95% 2-5 ml. Acetate Buffer pH 47 . . . . 97-5 ml. Methyl green, OO is washed repeatedly with chloroform, to extract all traces of methyl violet. The washed methyl green is then dried, and 0-15 gm. is weighed out and dissolved in the acetate buffer and the alcohol with the pyronin. Note: the stain is liable to deteriorate after two or three weeks. * See note {d) at the end of this technique. 3. fRibonuclease o-i% in distilled water which has been adjusted to pH 6-o t If this is not available a suitable ribonuclease extract can be prepared in the laboratory as follows: I. Pass 0-5 to I kilo Ox pancreas through a meat mincer. II. Pound it to a smooth pulp with a mortar and pestle. III. Suspend the pulp in one or two volumes of N/io acetic acid for sixteen to twenty-four hours. IV. Boil for ten minutes. V. Cool; then filter. VI. Adjust the pH of the filtrate to pH 6-o. VII. Filter. VIII. Add a crystal of thymol or camphor, as a preservative and store in the refrigerator: under these conditions the solution will keep for several months. Technique: Note: It is not possible to give absolute rules concerning the following procedure, as the methods of fixation, the solubility of the ribonucleic acid of the organ to be studied, and the activity of the ribonuclease, are all possible variables. The technique given below, which should be regarded as a basis for experiment, may be varied to suit individual lines of research and investigation. I . Fresh material should be used and objects or slices of tissue, which must not be more than 2-3 mm. in thickness, are fixed for a maximum of one hour in one of the above fixatives. 2a. After Zenker, wash with tap water for twenty-four hours. 2h. After Carnoy or Serra, wash in two or three changes of 95% alcohol for the minimum time necessary to remove the fixative. 3. Dehydrate and embed at once, through alcohol and toluol, in paraffin wax. 337 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES 4. Fix sections alternatively to slides marked A, B and C. 5. Remove paraffin wax with toluol, and pass through the usual descending grades of alcohol to distilled water. 6. Stain slide A immediately with the methyl green-pyronin for twenty minutes. 7. Meanwhile place slide B in distilled water, adjusted to pH 6-0 in the oven and leave there for an hour at 37° C. 8. Place slide C in the ribonuclease solution in the oven at 37° C. and leave there for an hour. 9. Meanwhile take slide A, after it has been in the methyl green-pyronin for twenty minutes, and wash it rapidly with dis- tilled water. 10. Slide A is then differentiated with 95% alcohol for five to ten minutes. 11. Dehydrate slide A with absolute alcohol; then wash with toluol and mount in balsam or D.P.X. or Cristalite. 12. When slide B has been in the oven for an hour, stain it with the methyl green-pyronin, differentiate, dehydrate and mount exactly as in the case of slide A. 13. When slide C has been in the oven for an hour, stain, differentiate, dehydrate and mount exactly as slides A and B. 14. Examine and compare specimen C with A and B. Notes : (a) Methyl green is unique in that it will stain some, but not all basophilic substances. It will stain high-polymer DNA as it occurs in the nuclei, but it will not stain depolymerized DNA or RNA in any form. In specimen B (above) it will be found that although the methyl green staining may have been completely abolished, Feulgen's reaction remains unchanged. Interesting comparisons can be made between sets of slides as B and C (above) with sections from the pancreas, kidney, lung, liver, genital glands, skin, etc. (See Professor J. Brachet's original 1942 paper*). {h) It may be found necessary, depending upon the fixative used and the organ to be studied, to raise the concentration of the ribonuclease solution to as much as o-6% to obtain optimum results. 338 SECTION SIX (c) The quality of the dyes, methyl green and pyronin, is a factor of great importance if good results are to be obtained with this technique. (d) In Professor Brachet's original paper (1942) the Unna Pappenheim Methyl Green-pyronin, formula as below, was em- ployed : Methyl green, 00 0-15 gm. Pyronin » • « .. 0-25 gm Alcohol 96% 1 • . . 2*5 ml. Glycerin « . . 20 ml. Phenol 0*5% aqueous 1 • . . 100 ml. This stain requires very rapid differentiation with the 95% alcohol as the stain, particularly the pyronin is liable to be washed out completely: differentiation here requires a great deal of practice and skill. For this reason Professor Brachet (1953) recommends the modified formula. However, chloroform washed methyl green is somewhat troublesome to prepare, and many workers may prefer to use the original Unna Pappenheim Methyl Green- Pyronin. Results: Specimen A : The presence of ribonucleic acid (of the nucleoli and of the cytoplasm) is indicated as red particles, and the desoxyribonucleic acid (of the chromatin) of the same cell, as blue. Specimen B: Treatment with hot water abolishes the methyl green staining of desoxyribonucleic (while Feulgen reaction remains unchanged). The absence of the methyl green staining confirms the presence and locali'i^ation of desoxyribonucleic acid. Specimen C: Ribonuclease brings about a complete loss of basophilia due to ribonucleic acid, not basophilia due to mucopolysaccharides. The absence of the red granules indicates the presence and localiza- tion of ribonucleic acid. The Feulgen reaction of the nuclei is unchanged. 339 I ml. 2gm. MEDICAL AND BIOLOGICAL STAINING TECHNIQUES References : * Brachet, J. (1942), Archives de Biol., 53, 207-57. " Localisation des acides pentosenucleiques dans les tissues animaux et les oeufs d'Amphibiens en voie de developpement. " Brachet, J. (1953), Q. jf. Micr. Sc, 94, i-io. " The use of basic dyes and ribonuclease for the cytochemical detection of ribonucleic acid. " NILE BLUE SULPHATE - SAFRANIN An histochemical technique for demonstrating phospho- lipids in frozen sections Solutions required: A. Formalin 10% . . . . . . 100 ml. Calcium Chloride 1% Calcium Carbonate . . Shake well: filter before use. B. Safranin 1% aqueous . . . . 100 ml. Aniline Oil . . . . . . . . 2 drops C. Nile blue sulphate, saturated aqueous 100 ml. Sulphuric acid 0*5% . . . . 10 ml. Boil for 2 hours under reflux condenser. Filter before use. D. Acetic acid 5% aqueous. E. HCl cone. . . . . . . . . 0-5 ml. Distilled water . . . . . . 99-5 ml. Technique: 1. Fix material in solution A: then cut frozen sections, without embedding: or the material may be embedded, if desired, in gelatine or carbowax. Note: Frozen sections should not be stored in water for more than ten to fifteen minutes. 2. Stain in the safranin solution for five minutes. 3. Rinse in distilled water. 4. Stain in the nile blue sulphate for ninety minutes at 60° C. 5. Rinse in distilled water. 6. Immerse in acetone heated to 50° C. on a water bath. 7. Remove the acetone from the source of heat and allow the sections to remain in it for half an hour. 340 SECTION SIX 8. Differentiate in 5% acetic acid for thirty minutes. 9. Rinse thoroughly in distilled water. 10. Refine the differentiation in the 0-5% HCl (Solution D). 1 1 . Wash in several changes of distilled water. 12. Mount in glycerine jelly. Results: Phospholipids, blue. Nuclei, red. Reference: Menschik, Z. (1953), Stain Tech., 28, 13-18. PHOSPHOMOLYBDIC ACID - EOSIN For the histochemical localization of choline-containing lipids, in frozen sections Solutions required: A. Acetone . . . . . . . . i volume Ether . . * . . . . . i volume B. Chloroform . . . . . . . . i volume Absolute alcohol , . . . . . i volume C. Phosphomolybdic acid. I % in solution B. D. Stannous chloride 1%. in 3N hydrochloric acid. E. Eosin 1% aqueous. Technique: 1. Dip frozen sections into acetone-ether. 2. Immerse in the phosphomolybdic acid solution for fifteen minutes. 3. Rinse in solution B. 4. Dip into solution D. 5. Counterstain with the eosin solution for one to two minutes. 6. Mount in glycerine jelly. Results: Positive areas are stained blue, whilst negative areas are red. Reference: Landing, B. H. (1952), Uzman, L. L. and Whipple, Ann. Lab. Invest., I, 456-2. MEDICAL AND BIOLOGICAL STAINING TECHNIQUES SUDAN BLACK The use of Sudan Black, also known as Sudan Black B, has been described in various works on Histochemistry. As far as I am aware, the correct structural formula of the Sudan Black molecule has not yet been disclosed in any literature published in any country, although it appears that this information has fre- quently been sought after by medical and biological research workers. It may, therefore, be of interest and help to research workers to give the formula here : H ^^N = N-<;^_^N = N-<^_^N\ yCR \ / \ >~N/ \CH. I H The molecular weight and other information regarding Sudan Black will be found on page 435. 34^ SECTION 7-SMEAR PREPARATIONS 2A ALBERTS STAIN (Laybourn's modification) For diphtheria bacilli Solutions required: A. Toluidine Blue .. 0-15 gm Malachite Green . . . . 0-2 gm. 95% alcohol . . 2 ml. Glacial acetic acid I ml. Distilled water . . 100 ml. 11* 11 Dissolve by warming or shaking well; allow to stand for twenty-four hours ; then filter. B. Potassium iodide . . • • 3 gm. Iodine . . . . . . . . 2 gm. Distilled water . . . . . . 300 ml. Dissolve the potass, iodide in about 5 ml. of the distilled water; add the iodine and shake until dis- solved ; then add the remainder of the distilled water and shake well. Technique: 1 . Fix films by gentle heat ; then stain for five minutes with Solution A. 2. Without washing apply Solution B for one minute. 3. Wash rapidly with water; blot dry. Results: Granules of diphtheria bacilli are stained bluish-black ; bars of bacilli dark green to black. Body of cells, light green; other organisms, light green. Note: Best results are obtained with a young culture (eighteen to twenty-four hours old) on serum medium. 345 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES ALCIAN BLUE For bacterial polysaccharides and capsules Note: This dye will stain bacterial capsules and insoluble polysac- charides in both pure and mixed cultures of bacteria and protozoa. It is specific for bacterial polysaccharides, and enzymatic degradation of these results in the loss of this stain. Solutions required: A. Alcian blue . . . . . . . . i gm. Alcohol 95% ,. .. .. 100 ml. B. Solution A . . . . . . . . i volume Distilled water . . . . • • 9 volumes Note: This solution which deteriorates within a few days, should only be prepared as and when required for immediate use. C. Carbol fuchsin (Ziehl-Neelsen) Technique: 1 . Fix air-dried smears by flaming in the usual way. 2. Stain with solution B for one minute. 3. Pour off excess stain and wash with water; then allow the preparation to dry in the air. 4. Counterstain with carbol fuchsin for a few seconds. 5. Wash immediately with distilled water to prevent over- staining. 6. Allow the preparation to dry in the air ; then examine. Results: Capsule and other bacterial polysaccharides are stained blue: cellular material, red. Note: Various carbohydrates including adonitol, arabinose, cello- biose, dextrin, glucose, inositol, inulin, mannitol, mannose, rhamnose, sorbitol, trehalose, xylose, and certain enzymes including papain, pepsin, rennin gave a positive reaction with alcian blue. The fact that the internal polysaccharides of the cell remain unstained is attributed to the complexity of the dye molecule which prevents its penetration of the cell wall. Reference: McKinney, Ross E. (1953), J^. Bact., (U.S.A.), 66, 453-4, ** Stain- ing bacterial polysaccharides. " SECTION SEVEN ANILINE BLUE - EOSIN B A simple and rapid technique for spermatozoa, which is particularly suitable for dog and human semen Solutions required: A. Ether I volume Absolute alcohol I volume B. Aniline blue aqueous . . 2 gm. Eosin, B . . I gm. Phenol I % aqueous . . 20 ml. Distilled water 60 ml. Technique: 1. Fresh semen is allowed to stand for about one half to one hour until it liquifies. 2. Prepare thin even smears of the liquified semen on scrupul- ously clean and dry slides or coverslips. 3. Fix for 3 minutes in a mixture consisting of equal volumes of ether and absolute alcohol ; then allow to dry in the air. 4. Flood smears with solution B and warm over a steam bath or hot plate while the stain is allowed to act for 5-7 minutes at 40-60° C. 5. Pour off excess stain. 6. Wash the preparation thoroughly with distilled water. 7. Drain and allow to dry thoroughly in air. 8. Mount in a neutral synthetic resin such as D.P.X., Clear- mount or Cristallite. Results: Sperm structure Stain reaction Galea capitis Pale bluish-grey; sharply outlined Cell membrane Bluish-grey; sharply outlined Nuclear membrane (shell) Bluish-grey ; sharply outlined Acrosome Slate blue Nucleas Pink Neck Sharply outlined (dark blue), inside colourless 347 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES End knobs Dark blue Middle piece Sheath dark blue, centre dark pink Axial filament Dark blue Tail Dark blue Notes: It is suggested that the technique which has been tried out only on dog and human semen, might be applied to other species. The technique gives good differentiation and preservation of cytological structure, reliable fixation and staining and undistorted and easily recognizable detail, upon which assays of semen for fertility depend. There is a tendency for abnormal forms of spermatozoa to stain more intensely than the normal forms. Reference: Casarett, George W. (1953), Stain Tech., 28, no. 3, 125-7. ANILINE GENTIAN VIOLET A simple and rapid stain for Treponema pallida Solutions required: A. Aniline gentian violet. B. Sodium hydroxide 1% aqueous 50 ml. Absolute alcohol ., .. 1-5 ml. C. Alcohol 5%. Technique: 1 . A loopful of serum from the lesion is spread into a film on a slide and dried in air. If an enlarged syphilitic gland is to be examined, 0-5 ml. sterile saline should be injected into the gland; then aspirated with a i-ml. syringe with a 22-gauge needle. Haemoglobin may be removed, if necessary, with distilled water. 2. Stain with ten drops each of Solutions A, B and C for two minutes, rocking the slide to ensure thorough mixing. 3. Wash in running tap water for twenty seconds; dry and examine. 348 SECTION SEVEN Results: Treponema pallida, deep purple, with distinct regular spirals free from precipitate. Spirochaeta refringens and other spiro- chaetes, purplish black, with regular open and coarse spirals. AZUR L For the detection and staining of epidermophytic infection Solution required: Azur L 0-5% aqueous. Technique: 1. Scrapings from around the margin of non-purulent areas of infected skin or from the tops of vesicles are transferred to a slide. 2. Treat three to ten minutes, depending on the size and thick- ness of the specimen, with Carnoy's fluid; then pour off carefully so that any floating debris is carried away and the fungus is left adhering to the slide. 3. Dry over a flame, taking care that the remaining Carnoy's fluid does not take fire. 4. Stain two or three minutes with 0*5% aqueous Azur L. 5. Rinse by adding distilled water drop by drop, taking care to avoid washing away the specimen. 6. Drain well ; then dry cautiously over a flame and mount. Results: Fungal filaments, dark blue against a light blue background. BASIC FUCHSIN For Treponema pallida in smears Solutions required: A. Potassium hydroxide 1% aqueous. B. Basic fuchsin 10% in absolute alcohol . . . . . . • • 5 rnl- Distilled water . . . . • • 95 ^• 349 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES Technique : 1. Fresh, air-dried smears are fixed for five minutes in pure methyl alcohol. 2. Allow the smears to dry in air ; then add a few drops of i % potassium hydroxide aqueous solution followed immediately by twice the number of drops of basic fuchsin solution. 3. Rock the slide to ensure thorough mixing of the two solu- tions; then leave until the liquid becomes decolorized, which should take about three minutes. 4. Pour off excess and rinse the preparation thoroughly with distilled water. 5. Drain, and blot dry carefully but thoroughly. 6. If it is desired to preserve the preparation it should be mounted at once in Cristalite. Results: Treponema pallidum, scarlet. Background, pink. BREED'S STAIN For staining and counting bacteria in milk Solution required: Methylene Blue . . . . . . 0-3 gm. 95% alcohol .. .. .. 30 ml. Distilled water . . . . . . 100 ml. Phenol . . . . . . . . 2| gm. Shake well until dissolved. Technique: 1 . Mark off i sq. cm. on a piece of white paper and superimpose a slide. 2. Place 00 1 ml. of the milk to be tested on the slide and spread, by means of a needle, into a film exactly i sq. cm., as indi- cated by the marking on the paper. 3. Dry on a level surface, by heating gently. 4. Immerse in xylol for a few minutes to remove the fat ; drain well; wash with absolute alcohol; then 95% alcohol; immerse in 90% alcohol for a few minutes. 350 SECTION SEVEN 5. Stain for two minutes with Breed's stain, prepared as above. 6. Wash quickly in 90-95% alcohol until the intense blue colour changes to a faint tinge. 7. Dry and examine. Results: Bacteria are stained dark blue against a lighter blue background BRILLIANT CRESYL BLUE For reticulated cells and platelets Solutions required: A. Brilliant Cresyl Blue 0-3% in pure absolute ethyl alcohol. B. Leishman stain or Wright's stain. Technique: 1. Place a drop of 0-3% Brilliant Cresyl Blue stain in absolute alcohol on a slide and allow it to dry. 2. A drop of blood 2 to 3 mm. in diameter is placed on another slide and brought in contact with the dried stain; the two slides are then manipulated hinge-like until all the stain has gone into solution and the blood appears blue-black. Allow the slides to come into contact to spread the drop ; then separate the slides and allow the films to dry. 3. Counterstain with Leishman or Wright by the standard technique. Results : Reticulum of immature red cells is stained clear cut blue; backgroiind, pale blue (fresh), or pink. Blood platelets, pale blue or lilac. Note: The counterstain may be omitted if it is desired only to count the platelets. The number of red cells per cm. should be determined separ- ately in a haemocytometer, and the ratio of platelets to red cells computed from the stained preparation. 351 MEDICAL AND BIOLOGICAL STAINING TECHNIQUES For reticulum Stock solution: Brilliant Cresyl Blue . . . ; i\ gm. Normal saline (0-85% NaCl) . . 100 ml. Technique: 1 . Mix a small quantity of the stock solution of the stain with 140 times its volume of normal saline solution. 2. Mix the blood in a white-cell counting pipette in the propor- tion of I volume blood to 20 vols, of the diluted staining solution. Shake the mixture for five minutes in the pipette, and place in a blood counting cell. 3. The fresh preparations are sealed with Vaseline to prevent drying, and are counted immediately. At least 1,000 should be counted f