2.0 CONNECTIVE TISSUES

2.1 VERHOEFF-VAN GIESON STAIN FOR ELASTIC FIBRES
Verhoeff's Technique (Verhoeff 1908) Elastic Fibres are composed of the protein, elastin. They are found widely distributed throughout the body but are especially associated with the respiratory system, circulatory system and the skin.

PRINCIPLE
The rationale of the technique is obscure; both iodine and ferric chloride with which the haematoxylin is combined, are oxidising agents and it is this property which probably accounts for the production of a black dye with cationic properties rather than a simple dye- mordant-tissue mechanism.

REFERENCES
p 32 Verhoeff's Technique, H C Cook Manual of Histological Demonstration Techniques. p 132 Verhoeff's Method for Elastic Fibres, Bancroft J D & Stevens A Theory and Practice of Histological Techniques, 4th Edition.

SPECIMEN
Standard paraffin section. This technique works after any fixative.

CONTROL
Dermis of skin, medium-sized arteries.

NOTES: Bulk staining is not possible due to the differentiation involved in this technique. Careful differentiation of each section is required, to ensure that the fine fibres are stained and at the same time a well-differentiated background is obtained. Elastic staining is extracted by the Van Gieson counterstain, therefore the section should be under-differentiated with ferric chloride. Pre-treatment with 1% potassium permanganate for 5 minutes, followed by oxalic acid, improves sharpness and intensity of staining.

REAGENTS
(A) 5 % ALCOHOLIC HAEMATOXYLIN
Haematoxylin 5.0g Absolute alcohol 100mL Dissolve the haematoxylin in approx 20mL of warm alcohol. Make up volume to 100 mL then filter. Leave to ripen naturally for one month before use.

(B) 10 % AQUEOUS FERRIC CHLORIDE
Take 16.5 mL commercial aqueous ferric chloride solution and make up to 100mL with distilled water.

(C) VERHOEFF IODINE SOLUTION
Dissolve 4 g of Potassium iodide in approx 10mL of distilled water. Add and dissolve 2 g of iodine in this solution and finally make up to 200mL with distilled water.

(D) VERHOEFF'S MIXTURE Working Solution To 5.0 mL of A add 2.0 mL of B, mix and add 2.0 mL of C, mix. May be used up to 48 hours after mixing.

(E) 2 % FERRIC CHLORIDE
Take 3.3 mL commercial 60% ferric chloride solution and make up to 100 mL with distilled water.

(F) VAN GIESON STAIN
Saturated Picric Acid 100mL 1 % aqueous Acid Fuchsin 10mL Boil for 3 mins, cool and filter, then add 0.25 mL concentrated Hydrochloric Acid.

PROCEDURE
1. Deparaffinise sections and hydrate to distilled water.
2. Stain in freshly prepared Verhoeff's mixture in a closed container for 15 mins.
3. Wash in water.
4. Differentiate in 2% Ferric chloride solution, checking each section microscopically until the nuclei and fine elastic fibres are black, but the background is still weakly stained.
5. Wash in water then in 95 % alcohol momentarily to remove iodine colouration.
6. Wash in water for 5 mins.
7. Counter stain with Van Gieson stain for 3 mins. Drain.
8. Dehydrate rapidly in alcohols - (Van Gieson stain will be removed if dehydration is prolonged).
9. Clear and mount.

RESULTS
Nuclei and elastic fibres Black
Collagen fibres Red
Background Yellow

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2.2 MASSON'S TRICHROME STAIN FOR COLLAGEN

Masson's Trichrome Method (1929) - Connective Tissue. The term "trichrome stain" is a general name for a number of techniques for the selective demonstration of muscle, collagen fibres, fibrin and erythrocytes. Three dyes are used, one of which may be a nuclear stain.

PRINCIPLE
Whilst the firm attachment of dye to tissue is probably electrostatic in nature, the means by which selective sequential staining with a number of anionic dyes is achieved is almost certainly bound up with physical structure of dye and tissue. The less porous tissues are coloured by the smallest dye molecule; whenever a dye of large molecular size is able to penetrate, it will always do so at the expense of the smaller molecule. Dyes, which are low in pH, tend to decolourise the nuclear stain, therefore iron haematoxylins which are more resistant to acid are used in preference to alum haematoxylins. Phosphomolybdic acid acts as a differentiator. Masson's method is a sensitive technique for displaying fine collagen fibres, basement membranes, fibrin and hyaline. Muscle is well shown in contrast and striations of voluntary muscle can be seen.

REFERENCE
p 94 Lee. G. Luna, Manual of Histologic Staining Methods of AFIP p 129 Bancroft and Stevens, Theory and Practice of Histological Techniques, 4th Ed. 1996.

SPECIMEN
Mercuric chloride or picric acid fixation gives best results. If tissue was fixed in neutral buffered formalin, staining is enhanced by using Bouin's fluid as a mordant. Standard paraffin sections.

CONTROL
Large artery: tunica media should stain red and the tunica adventitia blue or green. Section containing connective tissue, eg skin

REAGENTS
(A) BOUIN'S FIXATIVE.
Pre-heated to 56 - 60øC. Picric acid, saturated aqueous solution 750 mL 37% - 40% formalin 250 mL Glacial Acetic acid 50 mL
(B) WEIGERT'S HAEMATOXYLIN (IRON)
These solutions keep for approximately 6 months. a) Dissolve 1 g haematoxylin in 100 mL of 95% alcohol. Filter. Allow to ripen naturally for four weeks before use. b) Dilute 2 mL 60% commercial aqueous ferric chloride in 97 mL of distilled water. Add 1 mL concentrated hydrochloric acid. Working Solution Mix equal volumes of (A) and (B) prior to use. This mixture should be a violet black colour and must be discarded if it is brown.
(C) ACID FUCHSIN SOLUTION (kept in fridge) Acid fuchsin (CI 42685) 0.5 g Glacial acetic acid 5 mL Distilled water 100 mL
(D) PHOSPHOMOLYBDIC ACID SOLUTION Phosphomolybdic acid 1.0 g Distilled water 100 mL
(E) METHYL BLUE SOLUTION (kept in fridge) Methyl blue (CI 42780) 2.0 g Glacial acetic acid 2.5 mL Distilled water 100 mL

PROCEDURE
1. Deparaffinise sections and hydrate to distilled water.
2. Mordant in Bouin's Fixative for 1 (one) hour at 56øC in water bath (or overnight at room temperature).
3. Cool and wash in running water until yellow colour disappears.
4. Stain in Weigert's haematoxylin for 10 mins.
5. Wash well in tap water.
6. Stain in acid fuchsin solution for 5 minutes.
7. Rinse in distilled water.
8. Treat with phosphomolybdic acid solution for 5 minutes.
9. Drain.
10. Stain with methyl blue solution for 5 minutes.
11. Rinse in distilled water.
12. Treat with 1% acetic acid for 2 minutes.
13. Dehydrate through alcohols.
14. Clear and mount.

RESULTS
Nuclei Black
Cytoplasm, muscle and erythrocytes Red
Collagen Blue
Note: Light green counterstain may be substituted for methyl blue.
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2.3 GOMORI'S ONE-STEP TRICHROME STAIN

REFERENCE
Gomori GL. A rapid one-step trichrome. Amer J Clin Path. 1950;20:661 p131

SPECIMEN
Standard paraffin section of well-fixed tissue or alcohol-fixed smear.

REAGENTS
(A) BOUIN'S FIXATIVE SOLUTION
Preheated to 56 -60?C. Prepare as for method 2.2
(B) WEIGERT'S IRON HAEMATOXYLIN WORKING SOLUTION Prepare as for method 2.2
(C) GOMORI'S TRICHROME STAIN
Chromotrope 2R 0.6 g
Light green CI 42095 0.3 g
Glacial acetic acid 1.0 mL
Phosphotingstic acid 0.8 g
Distilled water 100 mL
(D) 1% GLACIAL ACETIC ACID SOLUTION
Alternatively, use acid-alcohol step on linear stainer.
PROCEDURE
1. Deparaffinise and hydrate to distilled water.
2. Place in Bouin's fixative in oven at 56øC for one hour.
3. Wash well in running water until sections are clear.
4. Stain in Weigert's iron haematoxylin working solution for 10 minutes.
5. Rinse in tap water.
6. Stain in Gomori's trichrome stain for 15 to 20 minutes.
7. Rinse in 1% acetic acid (or use acid-alcohol in linear stainer).
8. Rinse in distilled water.
9. Dehydrate and clear in 95% ethyl alcohol, absolute ethyl alcohol and histolene, 2 changes each
10. 2 minutes each.
11. Mount with safety-mount.

RESULTS
Muscle fibres red
Collagen green
Nuclei blue to green
Notes
1. Gomori's chromium haematoxylin may be substituted for the Weigert's iron haematoxylin at step 4.
2. Aniline blue can be substituted for the light green.
3. Slides may be left in Bouin's solution overnight at room temperature.
4. If sections are too red, differentiate in 100 mL of 1% acetic acid to which 0.7 g of phosphotungstic acid has been added.

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2.4 VAN GIESON TECHNIQUE FOR COLLAGEN

(Van Gieson 1889) The Van Gieson Technique is a "trichrome" stain for the demonstration of connective tissue. It is suitable for showing coarse collagen fibres, but not recommended for fine collagen fibres or muscle collagen comparisons.

PRINCIPLE
The mixture of the two acid dyes in acid solution compete for available linkages. Acid fuchsin being a larger molecular size is restricted to more permeable collagen fibres whereas the diffusible picric acid penetrates the compact muscle, cytoplasm and red cells.
REFERENCES
p 4 H C Cook, Manual of Histological Demonstration Techniques p 127 Bancroft JD & Stevens, Theory and Practice of Histological Techniques 4th ed

SPECIMEN
Standard paraffin sections Fixation - 10% neutral buffered formalin

CONTROL
Section containing connective tissue. eg skin

REAGENTS

(A) WEIGERTS IRON HAEMATOXYLIN
These solutions keep for approximately 6 months.
1. Dissolve 1 g haematoxylin in 100 mL of 95% alcohol. Filter. Allow to ripen naturally for four weeks before use.
2. Dilute 2 mL 60% commercial aqueous ferric chloride in 97 mL of distilled water. Add 1 mL concentrated hydrochloric acid. Working Solution Mix equal volumes of (1) and (2) prior to use. This mixture should be a violet black colour and must be discarded if it is brown.

(B) VAN GIESON'S STAIN
100mLs saturated aqueous Picric Acid 10 mLs 1% aqueous acid fuchsin (C.I. 42685) Boil for 3 minutes, cool and filter then add 0.25mL concentrated hydrochloric acid.

PROCEDURE
1. Deparaffinize and hydrate to distilled water.
2. Stain nuclei with Weigert's Iron Haematoxylin for 15 to 30 mins.
3. Wash in water until nuclei are dark blue-black and the background a paler blue-black.
4. Stain with Van Gieson's solution for 3 mins.
5. Dehydrate rapidly in alcohol.
6. Clear and mount.

NOTES
Acid fuchsin is extracted by water and picric acid by alcohol, hence rapid washing, blotting and brief dehydration are advised. Iron haematoxylin is used in preference to alum haematoxylin as it is more resistant to extraction by picric acid.

RESULTS
Nuclei Brown - black Collagen Red Muscle, RBCs, cytoplasm Yellow
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2.5 RETICULIN FIBRES

 PRINCIPLE
Reticulin fibres have little affinity for silver solutions and must be treated with a suitable solution, (2.5% iron alum), to produce sensitised sites on the fibres where silver deposition may be initiated. A separate reducing agent is required to cause deposition of silver in the form of the metal. Any unprecipitated, excess silver is removed by treatment with sodium thiosulphate solution.

CLINICAL SIGNIFICANCE
This method demonstrates the presence of reticulin in tissue sections.

REFERENCES
Bancroft, J.D. and Stevens, A.S, Theory and Practice of Histology Techniques. 2.110.1977.

SPECIMEN
A 5 micron paraffin section of tissue.

CONTROL
A section of normal liver tissue provides a good positive control.

REAGENTS
(A) Acidified Potassium Permanganate 0.5% Potassium permanganate 0.5g Distilled water 100mL 3% Sulphuric acid 5mL Mix immediately before use.
(B) 1% Oxalic acid
(C) 2.5% Iron alum (ferric ammonium sulphate).
(D) Ammoniacal silver 10% silver nitrate. 6mL Absolute alcohol 3mL Add strong ammonia dropwise, until precipitate that first forms, is just dissolved. (approx. 17 drops).
NOTE:
Use a conical flask to make up the silver solution and swirl it well in between each additional drop. This is important because the addition of too much ammonia renders the silver solution less effective. The silver solution should be made fresh each day it is required.
(E) Developer Formalin (40%) 10mL Absolute alcohol 20mL Distilled water 30mL

PROCEDURE
1. De-paraffinise and rehydrate to distilled water
2. Acidified potassium permanganate- 10 minutes..
3. Rinse off in distilled water
4. Oxalic acid - 3 minutes.
5. Rinse in distilled water
6. Iron alum - 15 minutes
7. Rinse with distilled water 3-4 times
8. This cuts down background staining.
9. Ammoniacal silver - 70 seconds.
10. For this step, best results are achieved by staining each slide separately. When applying the silver solution, pour on, tip off and re-apply all in one action. This ensures even staining of the section.
11. Wash x3 in distilled water - each wash 10 seconds.
12. Developer - about 3 minutes when it turns golden.
13. Wash well in water.
14. Fix in sodium thiosulphate (hypo). - 3 minutes
15. Wash well in water
16. Counterstain in 0.1% neutral red - 1 minute
17. Dehydrate, clear and mount.

RESULTS
Reticulin fibres - black
Tissue - red.

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2.6 GORDON & SWEETS' METHOD FOR RETICULIN FIBRES

(Gordon & Sweets 1936) Reticulin fibres are fine delicate fibres, which are normally found connected to stronger and coarser collagenic fibres. They make up the bulk of the supporting framework of the liver, spleen and lymph nodes.

PRINCIPLE
Silver from silver oxides is selectively deposited on the reticulin fibres, which appear black after conversion to reduced silver, by the reducing agent (formalin). Gold chloride is used as a toner to give a clearer background and unreduced silver is removed by treatment with Sodium thiosulphate.

REFERENCES
IMVS Method Manual 1986, Gordon & Sweets' Method for Reticular Fibres p 21 Cook H C, Manual of Histological Demonstration Techniques p 135 Bancroft JD & Stevens, Theory and Practice of Histological Techniques 4th Ed.

SPECIMEN
Standard paraffin sections Fixation 10% neutral buffered formalin

CONTROL
Liver/spleen

REAGENTS

(A) SILVER SOLUTION To 5mL of 10% Silver nitrate solution add strong ammonia until the precipitate formed is just dissolved. Avoid excess of ammonia. Add 5mL of 3% Sodium hydroxide solution. Re-dissolve the precipitate by adding strong ammonia drop by drop until the resultant solution retains a faint opalescence. If at this stage any excess ammonia is present, indicated by the absence of opalescence, add a few drops of 10% silver nitrate solution to produce a light precipitate. Make up to 50 mL with distilled water. Store in a dark bottle at 4øC - keeps 3 months. Filter before use (approximately 2 mL per slide). Note: All glassware should be rinsed in distilled water and dried before use.
(B) ACIDIFIED POTASSIUM PERMANGANATE
0.5% Potassium permanganate 190 mL 3% Sulphuric acid 10 mL (
C) 1 % OXALIC ACID
(D) 4 % IRON ALUM (Ammonium Ferric Sulphate)
(E) 10 % FORMALIN (not buffered) - dilute in tap water not distilled (ref Cook p21)
(F) 0.2 % GOLD CHLORIDE (Sodium Chloro-aurate)
(G) 5 % SODIUM THIOSULPHATE
(H) 1 % NEUTRAL RED (C.I. 50040)

PROCEDURE
(use PLASTIC forceps - not metal)
1. Deparaffinize and hydrate to distilled water
2. Acidified potassium permanganate 2 min
3. Wash in water
4. 1 % Oxalic acid 2 min
5. Wash well in distilled water
6. 4 % Iron alum 10 min
7. Wash in distilled water
8. Silver solution (filter 2mLs per slide) 1 min
9. Rinse in distilled water
10. 10 % Formalin 2 min
11. Wash in water followed by distilled water
12. 0.2 % Gold chloride 2 min
13. Rinse in distilled water
14. 5 % (Sodium thiosulphate) 2 min
15. Wash in water
16. Counterstain with 1 % Neutral Red 2 min
17. Dehydrate rapidly, clear and mount
Note: Dehydrate rapidly as the neutral Red counterstain is removed in alcohol.

RESULTS
Reticulin fibres Black
Nuclei Black
Background Red

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2.7 MALLORY'S PHOSPHOTUNGSTIC ACID HAEMATOXYLIN (PTAH) FOR MUSCLE STRIATIONS

The PTAH stain demonstrates many tissue structures, particularly fibrin, muscle striations, cilia and glial fibres, plus many CNS structures.

PRINCIPLE
The mechanism by which two-colour staining is achieved from a mixture of haematein and phosphotungstic acid is obscure. Garland and Gaer (1964) concluded that the blue colour produced was a metachromatic-type staining effect.

REFERENCES
p 13 Cook H C, Manual of Histological Demonstration Techniques p 117 Bancroft JD & Stevens Theory and Practice of Histological Techniques, 2ndEd. SPECIMEN Standard Paraffin sections Fixation -10% neutral buffered formalin

CONTROL
Skeletal (striated) muscle

REAGENTS
(A) PTAH SOLUTION
1. PTAH solution naturally oxidised .
Haematoxylin (C.I. 75290) 0.5 g Phosphotungstic acid 5.0 g Distilled water 500 mL The haematoxylin is dissolved in 100mL of the distilled water, and the phosphotungstic acid in the remaining 400mL. The two solutions are then mixed. The stain is allowed to ripen naturally in a loosely stoppered bottle at room temperature in the light for several months. This solution is stable and useable for many years. or
2. PTAH Solution using Haematein Haematein 0.5 g Phosphotungstic acid 5.0 g Distilled water 500 mL The solids are dissolved in separate portions of the distilled water as above, then mixed. Store in airtight bottle. The stain is ready for use in 24 hours and does not need to be chemically ripened due to the use of Haematein. or
3. PTAH Solution chemically oxidised Haematoxylin (C.I. 75290) 0.5 g Phosphotungstic acid 10.0 g Distilled water 500 mL 0.25 % Aqueous potassium permanganate 25 mL The solids are dissolved in separate portions of distilled water as above, then mixed. The potassium permanganate is then added to chemically ripen the stain, which can be used in 24 hours, but peak staining activity is reached in 7 days. Continuing oxidation means that this stain has a comparatively short life.

(B) 0.25% AQUEOUS POTASSIUM PERMANGANATE

(C) 5% AQUEOUS OXALIC ACID

PROCEDURE
1. Deparaffinize and hydrate to distilled water
2. Oxidise in potassium permanganate for 5 min
3. Wash in water
4. Bleach in Oxalic acid until white
5. Wash in water
6. Filter and stain in PTAH overnight at room temperature
7. Drain off excess stain
8. Dehydrate rapidly
9. Clear and mount

NOTE:
Dehydration should be rapid since water and alcohol may remove the stain. Dehydration may be commenced in 95% alcohol, and CNS sections may require thorough washing in 95% alcohol for several minutes to remove excess red stain.

RESULTS
Muscle striations, neuroglia fibres Dark blue
Fibrin, amoebae Dark blue
Nuclei, cilia, red blood cells Blue
Myelin Lighter blue
Collagen, osteoid, cartilage, elastic fibres Deep brownish/red
Cytoplasm Pale pinkish brown
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2.8 SIRIUS RED FOR COLLAGEN AND BILE PIGMENTS IN LIVER TISSUE.

The Sirius Red technique is a useful stain for collagen and bile pigment in liver biopsies.

PRINCIPLE
Collagen forms the ground substance of connective tissue. It is composed of three amino- acids and stains strongly with acid red dyes due to the affinity of the cationic groups of the proteins for the anionic reactive groups of the acid dyes.

REFERENCES
1. P174 Armed Forces Institute of Pathology, 3rd Edition.
2. Fouchet A (1917), Methode nouvelle de recherche et de dosage des pigments biliares dans le serum sanguin.Comptes rendus des seances de la Societe de la Societe de biologie et de ses filiales 80, 826.
3. Personal communication from Dr Pauline Hall, Specialist Pathologist, Flinders Medical Centre, Adelaide 3/8/95.

SPECIMEN
Standard paraffin section. Neutral buffered formalin fixation.

CONTROL
Normal liver tissue.

REAGENTS
(A) Fouchet's Reagent:
Trichloracetic acid 25 g
Distilled water 100 mL
10% Ferric chloride 10 mL
(B) Sirius Red (working solution):
Saturated aqueous Picric acid 98 mL
1% Aqueous Sirius Red 2 mL (Direct Red 80 C.I. 35780)
(C) Celestine Blue:
Celestine Blue (C.I. 51050) 0.5 g
Iron Alum (Ammonium Ferric Sulphate) 5 g
Distilled water 100 mL

PROCEDURE
1. Deparaffinize and hydrate test and control sections to distilled water.
2. Fouchet's reagent for 5 minutes.
3. Rinse in distilled water.
4. Celestine blue for 2 minutes.
5. Rinse in distilled water.
6. Alum Haematoxylin for 3 minutes - use linear stainer.
7. Wash, - do NOT differentiate- blue in Lithium Carbonate, wash.
8. Sirius Red WORKING solution for 20 minutes.
9. Rinse in distilled water.
10. Dehydrate, clear and mount.

RESULTS
Bile products Emerald green
Red blood cells Light/pale green
Collagen Bright red
Hepatocytes Dull yellow
Nuclei Black

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