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Histology Stain Manual Open Educational Resource

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Histology Stain Manual Open Educational Resource DE MONTFORT UNIVERSITY School of Allied Health Sciences Histology Stain Manual Open Educational Resource http://www.val.biologycourses.co.uk Handbook updated last August 2009 Many of these recipes are freely available on the internet these days, and the ones in this book were used in student classes and have been tried and tested over the years. 1. The aim of this handbook……………………. 2. Introduction to this handbook…. 3. Major changes to previous versions….. 4. ALCIAN BLUE-PAS COMBINED TECHNIQUE….. 5. ALCIAN BLUE-CRITICAL ELECTROLYTE CONCENTRATIONS….. 6. ALDEHYDE-FUCHSINE 7 FEULGEN-SCHIFF'S REACTION for DEOXYRIBONUCLEIC ACID (DNA) 8. GIEMSA STAIN (for cytological/haematological smears)….. 9. GIEMSA STAIN (post-dichromate fixation)….. 10. GORDON AND SWEET’S RETICULIN METHOD….. 11. HAEMATOXYLIN AND EOSIN….. 12. LUXOL FAST BLUE (Normal Myelin/ Lipofuscins))…. 13. MARTIUS, SCARLET, BLUE (M.S.B.)…. 14. MASSON'S TRICHROME TECHNIQUE…. 15. MAYER’S HAEMATOXYLIN AND EOSIN…. 16. PALMGREN’S METHOD (modified) for NERVE FIBRES…. 17. PERIODIC ACID-SCHIFF (PAS) REACTION…. 18. SCOTT’S TAP WATER…. 19. SOUTHGATE’S MUCICARMINE METHOD…. 20. THE ROMANOWSKY STAINS (Romanowsky 1891)…. 21. TOLUIDINE BLUE… 22. VAN GIESON’S STAIN FOR COLLAGEN…. 23. Resources…. 2 1. The aim of this handbook……………………. This stain manual is an essential laboratory guide for biomedical science and similar degree programmes where students need to master a number of histochemical stains. All the stains have been used in student classes so are tried and tested, although as you probably appreciate, a number of alternate recipes are available and will be equally as good. 2. Introduction to this handbook…. Histological staining techniques are fundamental to any research laboratory, whether it is in a hospital, pharmaceutical company or other industrial setting. Although quite an ancient art, being able to carry out a good quality stain is still an essential skill today, and one that De Montfort University graduates are recognised for. The histologist is able to view structural details of cells and tissues, important for diagnosing disease, monitoring the effects of treatment or answering numerous other research questions. Good staining requires the samples to be handled in an appropriate way, sectioned carefully, fixed in an appropriate agent, and then stained and mounted for examination. You will experience these techniques in the laboratory in several of your modules, and be assessed on the quality of your technical ability. 3. Changes in 2007…. This manual was modified in 2007 to refine and refocus the main histological and histochemical methods taught on the course. It is unfortunate that many of the more complex and varied staining methods employed in our original text were no longer being employed in NHS laboratories, and in many labs indeed these processes are now fully automated. This manual therefore comprises of the most essential stains and also those chosen illustrate some key principles of histochemistry. Happy staining! 3 4. ALCIAN BLUE-PAS COMBINED TECHNIQUE….. This is a most useful technique in that apart from distinguishing between acid mucins and neutral mucins, it also serves to demonstrate most mucins in the one preparation. This means in practice that a negative result (i.e. alcian blue negative and PAS negative) can be taken to mean that a given substance is unlikely to be a mucin. The rationale of the method is that by first treating with alcian blue the acid mucins will stain and thus be unable to react with the subsequent PAS. By following on with the PAS only neutral mucins and carbohydrates, such as glycogen, will stain red. Should a haematoxylin nuclear stain be used it is important to stain weakly, to prevent cytoplasmic staining acting as a potential source of confusion with the alcian blue. Mayer's haematoxylin solution is especially suitable for this. Fixation Formalin, mercury and other fixatives. Sections Paraffin sections. Solutions 1. 1% alcian blue in 3% acetic acid. (note: alcian blue is a copper phthalocyanin dye compound, and at low pH will stain acid -including sulphated- mucopolysaccharides by salt linkage with the acidic groups.) 2. 1% aqueous periodic acid. 3. Schiff's reagent. Technique 1. Dewax sections in dewaxing agent 2. Take sections down alcohols to purified water. 3. Treat with alcian blue solution (pH2.5 or pH0.5) for 5 minutes. 4. Wash well in purified water. 5. Treat with 1% periodic acid solution for 5 minutes. 6. Wash well in purified water. 7. Stain with Schiff's reagent for 8 minutes. 8. Rinse in Scott's tap water and develop in fresh Scott’s tap water for 5 minutes until section turns to a red/magenta colour. 9. Dehydrate from 70% isopropanol up to absolute. 10. Clear in clearing agent and mount in DPX. Results Acid mucins…………………………………..blue Neutral mucins……………………………….red/magenta Mucin mixtures……………………………….purple 4 Nuclei………………………………………….pale blue 5. ALCIAN BLUE-CRITICAL ELECTROLYTE CONCENTRATIONS….. An electrolyte such as magnesium chloride can be used to inhibit alcian blue staining. The electrolyte concentration that successfully competes with the alcian blue to prevent staining is referred to as the critical electrolyte concentration (CEC). Various acid mucins have different CEC points, influenced by their molecular weights that can be identified by the use of increasing molarities of magnesium chloride in a buffered alcian blue solution. CEC Solutions Alcian blue 0.05 g Acetate buffer 0.2M pH 5.8 100 cm3 MgC12.6H2O (MWt = 203.3): added to prepare the molarity series (0.06, 0.2, 0.5, 0.7, 0.9 M) Technique 1. Dewax in dewaxing agent. 2. Take sections down alcohols to purified water. 3. Stain replicate sections with the series of alcian blue/MgCl2 molarities overnight at room temperature. 4. Wash well in purified water. 5. Dehydrate from 70% isopropanol, clear in clearing agent and mount in DPX. Results The following acid mucins stain blue at the various molarities 0.006M…………………………………………..all acid mucins 0.2-0.3M…………………………………………only weakly and strongly sulphated mucins 0.5-0.6M………………………… ……only strongly sulphated mucins 0.7-0.8M…………………………………………only heparin/heparan sulphate and keratan sulphate 0.9M…………………………………...only keratan sulphate 5 6. ALDEHYDE-FUCHSINE Aldehyde-fuchsine has a higher affinity for sulphated mucins and can thus be used to distinguish sulphated from carboxylated mucins. In the aldehyde-fuchsine staining technique a pre-oxidation stage (Lugol’s iodine) is used to expose the sulphur-containing moieties in the tissue. However, here oxidation is not required, though a longer staining time in the aldehyde-fuchsine is used. Technique 1. Take sections to water and rinse in 70% isopropanol. 2. Place in a Coplin jar of aldehyde-fuchsine for 20 minutes. 3. Wash in 95% isopropanol, followed by de-ionised water. 4. Dehydrate in absolute isopropanol (95% - 100%), clear and mount. Results Sulphated mucins ……….purple Carboxylated mucins…….blue 6 7 FEULGEN-SCHIFF'S REACTION for DEOXYRIBONUCLEIC ACID (DNA) This is used in carbohydrate histochemistry and stains for sugars attached to DNA (it is therefore an indirect method of detecting DNA). As it splits the sugars from the DNA molecule, Schiff’s reagent stains and detects them. The reaction is based upon the liberation of active aldehyde groups by breaking the purine-deoxyribose bond with acid hydrolysis. The aldehydes recolour Schiff’s reagent (leucofuchsin) giving a purple colour to nuclear chromatin. Fixation Most fixatives, except Bouin's. If your section is fixed in Zenker’s, follow the directions below. Sections Paraffin sections: - These must be well dried on the slides and should be attached with adhesive to avoid section loss. Solution Schiff's Reagent (see PAS sheet) Technique 1. Dewax sections and dehydrate down alcohols to water. 2. Rinse briefly in cold (room temp.) 3. Transfer to M HCI and transfer to M HCI at 60°C for an optimal time. Place a control section in purified water at 60°C for the same time. 4. Place in Schiff's reagent for 30-60 minutes. 5. Rinse in Scott's tap water and develop in fresh Scott’s tap water for 5 minutes until section turns to a red/magenta colour. 6. Rinse 3 times in 5% aqueous sodium thiosulphate. 7. Wash in purified water. 8. Dehydrate from 70% isopropanol, clear in clearing agent and mount in DPX. Results DNA ………………………………………………….magenta Cytoplasm ……………………………………………variable if counterstain used If Zenker’s – fixed. 1. Rinse in Lugol’s iodine for 2 minutes. 2. Decolourise in 5% Na thiosulphate for 2 minutes. 7 3. Rinse in water. 8. GIEMSA STAIN (for cytological/haematological smears)….. Fixation Any Sections Cytological/haematological smears and fine needle aspirates. Solutions Stock Giemsa stain (International Committee for Standardization in Haematology; Boon & Drijvar, 1986). Azure B-thiocyanate Dimethylsulphoxide (DMSO) Eosin Y disodium salt (CI) Methanol Dissolve the azure B in the DMSO and the eosin Y in the methanol, then mix the two solutions. Store in a dark bottle at room temperature. The life of the solution is prolonged by the addition of a few drops of 1.OM HCl to give an apparent pH 4.0 (N.B. this is not an aqueous solution!). (Lillie modification) also available commercially as a reagent ready for use. Azure A-eosinate 0.5 g Azure B-eosinate 2.5 g Methylene blue-eosinate 2.0 g Methylene blue chloride 1.0 g Glycerol 375 cm3 Methanol 375 cm3 Mix the methanol and glycerol, and dissolve the dyes in the mixture overnight. Complete dissolution by shaking the stain for 10 minutes. Store in a dark bottle at room temperature. Buffered Giemsa stain To 10cm3 of stock Giemsa stain add 90cm3 of pH 6.8 0.033M phosphate buffer (pH6.5 0.03M HEPES is preferable). Mix and filter before use. Prepare fresh for each set of smears. (For the azure B/eosin Y stain, ICSH recommend a 1 cm3 Giemsa to 15 cm3 buffer 8 dilution, with a staining time of 25 minutes for blood films, 35 minutes for bone marrow films, and no additional ROM stain.) Technique Use Coplin jars throughout.
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