Histology Stain Manual Open Educational Resource

DE MONTFORT UNIVERSITY School of Allied Health Sciences

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….

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!

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

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

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.

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.

1. Use methanol fixed, air-dried smears. 2. Stain with buffered Giemsa stain for 30 minutes (dilute stock solution 1:9 with Giemsa buffer). 3. Rinse in Giemsa (phosphate) buffer (pH 6.8) 2 minutes 4. Drain, air dry. (see technical note, step 4) 5. Clear clearing agent in mount in DPX.

Technical notes

The use of Coplin jars is advised to minimize the introduction of stain deposit to the slide surface. If this does happen, decolourize the preparation in methanol and proceed from Step 2.

Step 1: The slide and cell preparation must be absolutely dry to ensure an optimal result. Some methods combine Steps 1 and 2 (fix and stain together) but initial fixation in methanol is recommended.

Step 4: Drain the slide and set aside vertically in a rack to dry. Drying may be hastened by placing the slide in a current of air or on a hot plate at 40°C. The slide must be completely dry before clearing in clearing agent otherwise water droplets will be seen microscopically. ROM stains are soluble in alcohol, therefore slides stained by this method should not be allowed to come into contact with this chemical.

Results

Nuclei ...... purple Granules of eosinophils_ ...... bright red Granules of basophils ...... dark purple Granules of neutrophis ...... light purple/blue Platelets ...... pink to purple Chromatin

9. GIEMSA STAIN (post-dichromate fixation)…..

This is a classic staining method for the adrenal. After dichromate fixation the chromaffin granules stain a greenish-yellow colour with Romanowsky stains.

Fixation Any dichromate fixative

Sections Thin paraffin sections.

Technique

1. Use methanol fixed, air-dried smears. 2. Stain with buffered Giemsa stain for 10 minutes (dilute stock solution 1:9 with giemsa buffer). 3. Rinse in giemsa (phosphate) buffer (pH 6.8) 2 minutes 4. Drain, air dry. (see technical note, step 4) 5. Clear clearing agent in mount in DPX.

Results

Nuclei ...... purple Granules of eosinophils_ ...... bright red Granules of basophils ...... dark purple Granules of neutrophis ...... light purple/blue Platelets ...... pink to purple Chromatin

10. GORDON AND SWEET’S RETICULIN METHOD…..

Fixation Not critical, formalin recommended.

Sections Thin paraffin sections. An adhesive is advisable.

Solutions 5. 5% aqueous oxalic acid. 6. 2% aqueous ferric ammonium sulphate (iron alum). 7. Acidified potassium permanganate solution.

0.25% potassium permanganate 47.5 cm3 3% aqueous sulphuric acid 2.5 cm3

The above solutions can be kept as stock solutions which will keep for several weeks.

4. Ammoniacal silver solution. To a 5 cm3 of 10% aqueous silver nitrate add concentrated ammonia drop by drop with frequent mixing, until the formed precipitate just dissolves. Then add 5 cm3 of 3.1% aqueous sodium hydroxide and mix. A precipitate will form which gradually dissolves upon the addition of ammonia, drop by drop as before. Stop when there are only a few precipitate granules remaining. Make up the final volume to 50 cm3.

Technique 1. Take sections to deionised water. 2. Treat with acidified potassium permanganate solution for 5 minutes (use 1% periodic acid in the practical and then go to step 4). 3. Wash sections in deionised water and bleach with oxalic acid solution for 1 minute. Wash well in tap water. 4. Rinse in deionised water then treat with iron alum solution for 2 minutes. 5. Wash well in several changes of deionised water. In the fume cupboard. 6. Treat with ammoniacal silver solution for 4-5 minutes with agitation. 7. Wash well in several changes of deionised water. 8. Reduce in 10% formalin in tap water for 30 – 60 seconds with agitation (chocolate brown colour). 9. Wash treat with 5% sodium thiosulphate for 10 minutes. 10. Wash dehydrate from 70% isopropanol, clear in xylene and mount in DPX.

Results Reticulin fibre – black Collagen – brown to yellow-brown if untoned, purple-grey if toned. Background – red counterstain of used, if not, clear.

11. HAEMATOXYLIN AND EOSIN…..

“Taking Sections to Water” This simply refers to the procedure in which a specimen is hydrated by immersing it in DECREASING concentrations of alcohol (therefore containing more water).

“Dehydrating Sections” This is the reverse process in which the specimen is dehydrated again by going through INCREASING concentrations of alcohol.

1. De-wax paraffin wax section in histoclear for approx. 10 minutes. 2. “Take sections to water” - hydrate sections: a) Place in 100% isopropanol for 2 minutes. b) Transfer to second jar of 100% isopropanol for 2 minutes. c) Place in 95% isopropanol for 2 minutes. d) Transfer to second jar of 95% isopropanol for 2 minutes. e) Place in 70% isopropanol for 2 minutes. f) Place in purifed water for 1 minute.

3. Stain in Haematoxylin (usually Mayer’s) for 6 minutes. 4. Rinse in purified water. 5. Differentiate by adding a drop of acid-alcohol (0.5% HCl in 70% ethanol), enough to cover the section. 6. Rinse immediately in purified water. 7. Place section into a beaker of tap water which has had Scott’s tap water added to it (10 drops of Scott’s to 50 ml of water). Leave section to “blue” for 5 minutes. 8. Rinse in purified water. 9. Stain in 1% Eosin for 1 minute. 10. Rinse in 95% isopropanol. 11. “Dehyrate sections”: a) Place in 95% isopropanol for 5 minutes. b) Place in first jar of 100% isopropanol for 5 minutes. c) Transfer to second jar of 100% isopropropanol for 5 minutes.

12. Place in clearing agent (histoclear) for 10-15 minutes. 13. Cover section with coverslip and mount in DPX. 14. Clearly label your slide. 15. Place on drying plate.

12. LUXOL FAST BLUE (Normal Myelin/ Lipofuscins))….

Although luxol fast blue is slow in its action and does not give an intense colouration with myelin of the PNS, it is easy to use. The luxol dyes are arylguanidinium salts of anionic chromogens. They are hydrophobic molecules and will thus stain lipids and hydrophobic domains of proteins. When the dye attaches to the substrate an arylguanidinium gegen-ion is released, leaving behind the coloured anion. To make the dye specific for myelin the staining is differentiated by treating with lithium carbonate, followed by 70% ethanol. The use of a cationic counterstain (cresyl violet) will not only bind to nuclei and Nissl substance but enhance the luxol fast blue staining by binding to this anion, changing it from green to blue.

Fixation

Formal-acetic-methanol.

Sections

Thick (25pm) paraffin, frozen, low-viscosity nitrocellulose (LVN) sections.

Solutions Staining Solution Luxol fast blue 1 g 95% Ethanol 1000 cm3 10% Acetic acid 5 cm3

Technique

1. Dewax sections and dehydrate to 95°C isopropanol 2. Stain in luxol fast blue staining solution for 2 hours (30 minutes will be sufficient for the sections used in the class) at 60°C, or overnight at 37°C. 3. Wash in 95% isopropanol followed by purified water. 4. Start the differentiation process by immersing the section in 0.005% lithium carbonate for 10 minutes. 5. Differentiate in 70% ethanol until grey and white matters are clearly distinguishable (30 to 60 seconds). 6. Wash well in purified water and examine microscopically. The nuclei should be decolourized; repeat stages 4 and 5 if necessary. 7. Counterstain in 0.1% cresyl violet in 1% acetic acid for 10 minutes at 37°C. 13 8. Wash in purified water and rinse in 70% isopropranol. 9. Differentiate the cresyl violet in 95% isopropanol, until only the nuclei and Nissl substance are stained purple. 10. Rinse rapidly in absolute isopropanol, clear in clearing agent and mount in DPX.

For lipofuscins: 1. Counterstain in 1% aqueous neutral red. 2. Rinse in purified water. 3. Dehydrate in absolute isopropanol, clear in clearing agent and mount in DPX.

Results

Myelin ...... blue Nuclei--- ...... purple (cresyl violet counterstain) Nissl substance ...... purple Some ceroid lipofuscins ...... blue Nuclei ...... red (neutral red counterstain)

14 13. MARTIUS, SCARLET, BLUE (M.S.B.)….

The MSB technique is a modification of the Masson trichrome procedure and is used to demonstrate fibrin, especially older deposits. The small molecule of Martius yellow (acid yellow 24), together with phosphotungstic acid in alcoholic solution, selectively stains red cells and early fibrin deposits. The phosphotungstic acid blocks the yellow staining of muscle, collagen and most connective tissue fibres. The medium sized molecules of brilliant crystal scarlet (acid red 44) stain the muscle and mature fibrin, with again the collagen fibres being prevented from taking up the stain by phosphotungstic acid. Final treatment with the large molecule, soluble blue (methyl blue/acid blue 93), stains the old fibrin deposits and collagen.

Fixation Most fixatives but see notes for Masson's.

Sections Thin paraffin sections.

Solutions

Martius yellow Martius yellow (acid yellow, C. 1. 10315) 0.5 g Phosphotungstic acid 2.0 g 95% Ethanol 100 cm3

Dissolve Martius yellow in the alcohol before adding the phosphotungstic acid.

Brilliant crystal scarlet Brilliant crystal scarlet (acid red 44, C.I. 16250) 1.0 g Glacial acetic acid 2.0 cm3 Purified water 100 cm3

Methyl blue Methyl blue (acid blue 93. C.I. 42780) 0.5 g Glacial acetic acid 1.0 cm3 Purified water 80 cm3

Glacial acetic acid (1 % vN) Phosphotungstic acid (1% w/v)

15 Technique

1. Dewax sections and take down alcohols to water. 2. Acidify with 0.1M HCl for 1 minute. 3. Stain with Weigert's iron haematoxylin (Weigert’s A & B mixed – 1:1 ratio) for 6 - 10 minutes. 4. Differentiate in acid alcohol (0.5% HCI in 70% isopropanol) 5 seconds. 5. Blue in Scott’s tap water for 5 minutes. 6. Rinse in purified water 7. Rinse in 95% isopropanol. 8. Stain with Martius yellow for 5 minutes. 9. Rinse in purified water. 10. Stain in Brilliant crystal scarlet for 10 minutes. 11. Rinse in purified water. 12. Treat with phosphotungstic acid until no red remains in the collagen 5-10 minutes. 13. Rinse in purified water. 14. Stain in Methyl blue until collagen is coloured 5 minutes. 15. Rinse in 1% acetic acid. 16. Dehydrate from 70% isopropanol upwards. 17. Clear in clearing agent and mount in DPX.

Results Nuclei ...... blue Erythrocytes ...... yellow Muscle ...... red Collagen ...... blue Fibrin (early deposit) ...... yellow Fibrin (mature deposit) ...... red Fibrin (very old deposit) ...... blue

Technical notes The staining is dependent on the size of the dye molecule and a number of satisfactory substitutes can be made. Martius yellow can be substituted for by the slightly larger lissamine fast yellow molecule, which is less easily displaced by the subsequent red dye. Brilliant crystal scarlet can be replaced by anionic red dyes, such as Ponceau de Xylidine or azofuchsine, and the methyl blue dye replaced by durazol blue, pontamine sky blue, fast green FCF or naphthalene black 108.

Stage 12: Although 4 minutes will achieve a selective staining, preference may require 10 minutes.

Stage 14: Examine at 2 minute intervals; excessive stain cannot easily be removed.

Reference Lendrum, A.C., Fraser, D.S., Slidders, W. and Henderson, R. (1962). Studies on the character and staining of fibrin. Journal of Clinical Pathology, 15: 401

16 14. MASSON'S TRICHROME TECHNIQUE…. The success of this method depends largely on the degree of differentiation of ponceau- acid fuchsine by phosphomolybdic acid. It is important to prolong differentiation until the connective tissue is almost unstained. The technique is useful for differentiating collagen from other connective tissue.

Fixation Zenker, Helly, Bouin and formal-sublimate are especially recommended. 10% formalin may be adequate but unembedded formalin-fixed material will benefit from appropriate secondary fixation. Formalin fixed sections may be mordanted for up to 3 hours in saturated alcoholic picric acid, containing 3% mercuric chloride, followed by thorough washing to remove picric acid staining.

Sections Thin paraffin sections. An adhesive is advisable.

Solutions 1. Cytoplasmic (plasma) stain 1% ponceau de xylidine (ponceau 2R) in 1% acetic acid 2 parts 1% acid fuchsine in 1% acetic acid 1 part 2. Differentiator and mordant 1% phosphomolybdic acid in purified water 3. Fibre stain 2% light green in 1% acetic acid

Technique 1. Dewax sections and take down alcohols to purified water 2. Pre-treat with 0.1M HCl for 1 minute 3. Stain nuclei with Weigert's iron haematoxylin (Weigert’s A & B mixed – 1:1 ratio) for 10 minutes. 4. Wash sections well in scott’s tap water. 5. Differentiiate nuclear stain with 0.5% HCI in 70% alcohol and blue in scott’s tap water for 5 minutes. 6. Wash well in purified water. 7. Stain in the red cytoplasmic stain for 5-10 minutes 8. Rinse in purified water. 9. Differentiate in 1 % phosphomolybdic acid until collagen is decolourized, with muscle, erythrocytes and fibrin remaining red. 5 - 10 minutes are required. 10. Rinse in purified water. 11. Counterstain in light green for 1 minute. 12. Wash well in 1% acetic acid for at least 1 minute. Go directly to 95% alcohol wash. 13. Dehydrate in 100% alcohol, clear in clearing agent and mount in DPX.

Results Collagen, some reticulin, amyloid and mucin………………green Muscle, RBCs, fibrin and some cytoplasmic granules …….red Nuclei ...... ……….black

17 15. MAYER’S HAEMATOXYLIN AND EOSIN….

1. Dewax and hydrate to water. 2. Stain in Mayer’s haemalum for 6 minutes. 3. Differentiate in acid alcohol briefly 2-3 seconds. 4. Immerse in Scotts tap water to turn blue (prepare by adding 3-4 drops of Scots to tap water in a 100ml beaker). 5. Stain in eosin for 60 – 90 seconds. 6. Transfer to a coplin jar labeled 95% alcohol rinse and rinse off excess stain. 7. Dehydrate through ascending alcohols. 8. Clear, mount, label and leave to dry.

16. PALMGREN’S METHOD (modified) for NERVE FIBRES….

This method stains peripheral nerves using silver. In this method it is necessary that the silver impregnation leaves non-nervous tissues, such as reticulin and collagen, unstained.

Fixation 10% Formalin even after long fixation periods. (Osmium tetroxide, chromic acid, potassium dichromate and mercuric chloride should be avoided).

Sections Semi-thick (8pm) paraffin sections.

Acid Formalin Formalin 25 cm3 Purified water 75 cm3 1% nitric acid 0.2 cm3

Silver Solution Silver nitrate 15 g Potassium nitrate 10 g Purified water 100 cm3 5% glycine 1 cm3

The silver solution is much more stable than ammoniacal silver solutions and is still effective after 1 week, but best results are obtained using fresh solution.

Reducer Solution Plyrogallol 10 g Absolute ethanol 550 cm3 Purified water 450 cm3 1% nitric acid 0.2 cm3

The reducer keeps very well. It gradually darkens with age but remains usable for several months. It is important to remember, however that it can only be used once.

Technique

1. Dewax sections and take down alcohols to 95% isopropanol. 2. Place slide in 70% isopropanol for 2 minutes, then take to water. 3. Treat with acid formalin for at least 3 minutes. 4. Wash well in purified water. 5. Place in silver solution for 15 minutes at room temperature. 6. Drain slide of silver solution but do not rinse; transfer to preheated reducer solution for 1 minute at 40 to 45°C, agitating gently. Use a fresh Coplin jar of reducer for each section. 7. Wash well in purified water. 8. Fix in 5% sodium thiosulphate for 5 minutes. 19 9. Dehydrate through the isopropanols. 10. Clear in clearing agent, without agitating the slide, and mount in DPX. Results

Nerve fibres ...... ……….. dark brown to black Background …………………………………..golden brown

Additional information: Treatments with acid formalin, silver stains and reducer reagents will be located in the fume cupboard. 9. Dewax and hydrate to water. 10. When requested give your section to the lab demonstrator (make sure it is clearly labeled). 11. Your section will be placed in acidified formalin for 3 minutes following a brief water rinsing step. 12. When you hand your section in note the time and return to your bench. Please go back to the demonstrator after 5 minutes to see the next step. 13. This entails a 15 minutes incubation in the Palmgren’s silver reagent. Return to your bench and note the time. 14. After the silver staining go back to the demonstrator which is 45OC in Palmgrens reducer. Bring with you a coplin jar or 100ml beaker of water to collect your section. 15. Following silver staining proceed with the remaining steps from step 9 above.

Step 9 is similar to photographic techniques. The reagent removes all undeveloped silver from the film surface. In this method treatment with 5% sodium thiosulphate solution removes all unbound silver from the slide. This takes 10 minutes. Following this, wash briefly in deionised water, dehydrate, clear and mount in DPX.

20 17. PERIODIC ACID-SCHIFF (PAS) REACTION….

Adjacent 1:2 glycol groups (CHOH-CHOH) are broken by periodic acid and converted into aldehydes (two CHO groups); these are demonstrated with the Schiff's reagent.

The most important PAS positive carbohydrates in tissues are polysaccharides (glycogen), neutral mucopolysaccharides, mucoproteins, glycoproteins and glycolipids. Acid mucopotysaccharides are only weakly positive or negative. The PAS reaction can be used to demonstrate many other normal or pathological tissue constituents, the most important of which are:

Fixation Most fixatives may be used. For labile polysaccharides, fixation must be immediate, and thin pieces of tissue are required.

Sections Paraffin, frozen, freeze-dried or cryostat, glycol-methacrylate sections.

Solutions Schiff's Reagent Boil 200 cm3 of purified water and add 1g of basic fuchsine. When dissolved, cool and filter. Bubble SO2 gas slowly through the solution, shaking occasionally until it becomes a clear transparent red colour. Stand the stoppered flask in a dark cupboard overnight. If the solution is pale straw coloured or colourless next morning it is ready for use. If some residual red colour remains, decolourize with 1g of activated charcoal; shake and filter. Store at 4°C and discard when a pink colour develops.

Technique

1. Dewax sections (N.B. a sufficient number for control and test) and hydrate down alcohols to water 2. Oxidize for 5 minutes in 1% aqueous periodic acid solution. 3. Rinse in purified water. 4. Place in Schiff's reagent for 8 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. Dehydrate from 70% isopropanol up to absolute. 7. Clear in clearing agent and mount in DPX.

Technique to distinguish between glycogen and acid mucins.

1. Dewax sections (N.B. a sufficient number for control and test) and hydrate down alcohols to water 2. Oxidize for 5 minutes in 1% aqueous periodic acid. 3. Incubate test section and one glycogen +ve control section in 1% diatase for 1 hour at 37oC. 4. Rinse in purified water. 5. Place in Schiff's reagent for 8 minutes. 6. 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. 7. Dehydrate from 70% isopropanol up to absolute. 8. Clear in clearing agent and mount in DPX.

This technique is useful for distinguishing the true presence of glycogen from other acid mucins.

Results

PAS positive substances …………………………………… red or magenta Nuclei…………………………………………………………… variable if counterstain used

The most important PAS positive carbohydrates in tissues are polysaccharides (glycogen), neutral mucopotysaccharides, mucoproteins, glycoproteins and glycolipids. Acid mucopolysaccharides are only weakly positive or may be negative. The PAS reaction can be used to demonstrate many other normal and pathological tissue constituents, the most important of which are:

Amyloid Epithelial mucins Lipochrome Basement Fungi pigmentsMucoid cells of MembranesCartilage Glycogen anteriorPancreatic pAugary Cellulose Hyaline membrane Starch zymogen gram/es Cerebrosides ofneonatal lung Thyroid colloid

22 18. SCOTT’S TAP WATER….

Add 3-4 drops of Scott’s to tap water in a 100 ml beaker.

19. SOUTHGATE’S MUCICARMINE METHOD….

This is one of the classical histological techniques and gives clear permanent preparations. It is purely empirical, and has tended to be overshadowed by other methods that give an indication of the chemical composition of the mucin, but it is still recommended for the demonstration of epithelial mucin.

Fixation Formal saline or most other fixatives,

Sections Paraffin sections

Staining Solution Place 1g of powdered carmine and 1g of dry aluminium hydroxide in a 500 cm3 flask. Add 100 cm3 of 50% ethanol, then 0.5g of anhydrous aluminium chloride, whilst shaking. Place in a boiling water bath and boil for 2-3 minutes. Cool and filter.

Technique

1. Dewax sections in dewaxing agent 2. Bring sections down alcohols to purified water. 3. Stain nuclei with Mayer’s haematoxylin for 6 minutes. 4. Differentiate in acid alcohol and blue in Scott's tap water for 5 minutes. 5. Stain for 1 hour in Southgates Mucicarmine. 6. Rinse in purified water (recommend 3 changes of purified water) 7. Dehydrate from 70% isopropanol, clear in clearing agent and mount in DPX.

Results

Epithelial mucin (including gastric mucin)……..red Nuclei ...... ………blue

20. THE ROMANOWSKY STAINS (Romanowsky 1891)….

Often regarded as a special stain in histology the Romanowsky (ROM) technique must be regarded routine in any cytology department engaged in non-gynaecological specimen preparation. The technique was developed as a haematological stain for blood smears, with the dyes becoming a most useful diagnostic aid in cytology. Many ROM formulae are commercially available either as a dry powder or reagent ready for use. They are neutral dyes in that they consist of a combination of basic thiazine dyes (methylene blue and/or azure B) and the acid stain eosin which should be dissolved in acetone-free methanol for use.

Fixation Any

Sections Cytological/haematological smears and fine needle aspirates.

Solutions

ROM stain of choice available commercially, ready for use. Preparation of powdered dye:

Jenner 3 g/l in methanol Leishman 1.5g/l in methanol May- 3 g/l in methanol Grunwald Place the crystalline powder in a mortar and add a small aliquot of methanol. With a pestle grind the dye until a near saturated solution is apparent. Decant the stain into a dark bottle. Repeat the grinding and decanting process with more methanol until no more dye is extracted from the crystals. Store at room temperature and allow a few days for ripening. Filter fresh aliquots each day before use.

Stock Giemsa stain (International Committee for Standardization in Haematology; Boon & Drijvar, 1986). Azure B-thiocyanate 1.0 g Dimethylsulphoxide (DMSO) 133 cm3 Eosin Y disodium salt (C.I. 0.33 g Methanol45380) 200 cm3

Azure A-eosinate 0.5 g

Azure B-eosinate 2.5 g Methylene blue-eosinate 2.0 g Methylene blue chloride 1.0 g 3 Glycerol 375 cm Methanol 375 cm3

(Lillie modification) also available commercially as a reagent ready for use. 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 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.

1. Fix air-dried smears in methanol 2 minutes. 2. ROM stain of choice for 2 to 5 minutes. 3. Buffered Giemsa stain for 6 minutes (dilute stock solution 1:9 with Giemsa buffer). 4. Rinse in Giemsa (phosphate) buffer (pH 6.8) 2 minutes 5. Drain, air dry. (see technical note, step 5) 6. Clear clearing agent in mount in DPX.

Results

Nuclei ...... purple Degenerate nuclei ...... pink 26 Cytoplasm ...... pink or blue Cytoplasm of lymphocytes ...... slatey blue Red cells ...... pink/red Granules of eosinophils_ ...... bright red Granules of basophils ...... dark purple Granules of neutrophis ...... light purple/blue Platelets ...... pink to purple Chromatin Chromatin of Plasmodium and Leishmania (protozoan parasites ) ...... red

The ROM stain is particularly recommended for body cavity fluids and fine needle aspirations where haematological disorders may be expected. It can be helpful in distinguishing between lymphomas and small cell undifferentiated carcinoma. It is also useful for detecting fungi and parasites in smears.

Technical notes

The use of Coplin jars is advised to minimize the introduction of stain deposit to the slide surface. If this does happen, decolourize the preparation in methanol and proceed from Step 2.

Step 1: The slide and cell preparation must be absolutely dry to ensure an optimal result. Some methods combine Steps 1 and 2 (fix and stain together) but initial fixation in methanol is recommended.

Step 2: ROM stains are used undiluted, must be filtered fresh each day, and must not be allowed to dry on the surface of the slide as precipitation occurs readily.

Step 3: Buffered Giemsa stain, when used secondarily to Jenner, Leishman or May- Grunwald stains, considerably enhances cytoplasmic staining.

Step 4: This step differentiates the stain. Blue dye is removed slowly to accentuate the eosin colouration. Take into account the drying of the slide in the overall buffer rinse time.

Step 5: Drain the slide and set aside vertically in a rack to dry. Drying may be hastened by placing the slide in a current of air or on a hot plate at 40°C. The slide must be completely dry before clearing in clearing agent otherwise water droplets will be seen microscopically. ROM stains are soluble in alcohol, therefore slides stained by this method should not be allowed to come into contact with this chemical.

27 21. TOLUIDINE BLUE…

1. Take section to water. 2. Stain for 3 minutes in 0.5% toluidine blue pH 4. 3. Dehydrate, clear and mount.

22. VAN GIESON’S STAIN FOR COLLAGEN….

The Van Gieson stain uses two anionic dyes at low pH (1.0 to 2.0); a small dye molecule (picric acid) penetrates all the molecular spaces of the tissue, but is more easily washed out than a larger dye molecule (acid fuchsine) which can only enter the collagen matrix.

Fixation A wide range of fixatives give good results; mercuric chloride is especially suitable

Sections Paraffin sections.

Solutions Van Gieson's Stain

Saturated aqueous picric acid 100 cm3 1 % acid fuchsine (Cl 42685) in purified water 5-10 cm3

The exact proportion of acid fuchsine used will depend largely on the quality of the dye sample, the optimum concentration being readily determined. The mixture keeps well.

Technique

1. Dewax sections in dewaxing agent 2. Take sections down alcohols to purified water. 3. Pre-treat with 0.1M HCl for 1 minute 4. Stain nuclei with Weigert's iron haematoxylin (Weigert’s A & B mixed – 1:1 ratio) for 5 minutes. 5. Wash sections well in water and differentiate in acid alcohol. 6. Blue in tap water made alkaline with 4 - 5 drops of Scott's tap water, followed by a rinse in purified water. 7. Stain in Van Gieson's stain for 5 minutes. 8. Proceed directly to 95% isopropanol. Do not wash in water, especially alkaline water, as this will extract the red stain. 9. Dehydrate in absolute isopropanol, clear in clearing agent and mount in DPX.

Results

Collagen…………………………………………………………. deep red Nuclei……………………………………………………………….brown-black to black Muscle, cytoplasm, erythrocytes, fibrin………………………..yellow

23. Resources….

School of Anatomy and Human Biology - The University of Western Australia Website entitled “Blue Histology”. Great on-line resource for looking up sections and stains. http://www.lab.anhb.uwa.edu.au/mb140/

Stevens, A. & Lowe, J. Human Histology. 3rd edition, Gower./Mosby, (2004) (or second- hand previous editions)

Young, B. & Heath, J.W. Wheater’s Functional Histology. 4th edition, Churchill-Livingstone (2000) (or second-hand previous editions)

University of Bristol searchable database of stain protocols and results. http://www.bristol.ac.uk/vetpath/cpl/histmeth.htm

On-line photo gallery of stains and protocols at Immunohistochemistry World. http://www.ihcworld.com/imagegallery/