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….
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
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.
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 gAzure 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 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.