Procedures of Histochemical

ALCIAN BLUE-PERIODIC ACID-SCHIFF FOR MUCINS

Procedure 1. Stain in Alcian blue for 15 minutes.

2. Rinse in water.

3. Treat with 1% Periodic acid for 5 minutes.

4. Rinse with distilled water.

5. Treat with Schiff's reagent for 5 minutes.

6. Rinse in distilled water.

7. Put in running water for 5 minutes.

8. Counterstain with Harris haematoxylin for 2 minutes, differentiate and blue.

9. Dehydrate, clear and mount.

Results: Acid mucopolysaccharide blue PAS positive substance, neutral mucin magenta nuclei blue

Solutions: 3% alcian blue in 3% aqueous acetic acid - pH 2.5

Schiff's reagent Refer to protocol PAS

ALCIAN BLUE FOR ACIDIC MUCINS

Procedure 1. B.S.D.W.

2. Stain in Alcian blue for 10 minutes.

3. Rinse thoroughly in water.

4. Counterstain with 1% neutral red for 1 minute.

5. Rinse briefly in water.


Results: Acid mucopolysaccharide blue

pH 3.1 general acidic mucins

pH 2.5 carboxylated and sulphated mucins

pH 1 sulphated mucins only

pH 0.2 strongly sulphated mucins only

Solutions: pH 3. 1 Dissolve 1 g alcian blue in 100 ml 0.5% acetic acid

pH 2.5 Dissolve 1 g alcian blue in 100 ml 3.0% acetic acid

pH 1.0 Dissolve 1 g alcian blue in 100 ml N/10 HCl

pH 0.2 Dissolve 1 g alcian blue in 100 ml 10% sulphuric acid

Notes: Hyaluronidase digestion 1 mg type IV testicular hyaluronidase in 1 ml pH 6.7 phosphate

buffer for 3 hours at 37 °C (preheat before use)

KERATIN - MUCIN

Procedure: 1. Stain section with filtered alcian blue solution for 10 minutes.

2. Rinse with water.

3. Stain nuclei with Harris' Hx, differentiate and blue.

4. Stain with phloxine solution for 5 - 15 minutes, blot dry.

5. Differentiate in tartrazine until a yellow background with red keratin and RBC.

6. Rinse with 95% alcohol, rinse in abs. alcohol, clear in xylene and mount.

Results: Mucin Blue to green Keratin, RBC, Fibrin Red

Nuclei Bluish grey

Other structures Yellow

Solutions: Alcian blue solution:

o 1% Aq. alcian blue 100 ml

o 1% Aq. acetic acid 100 ml

Phloxine solution:

o Phloxine 0.5 gm Calcium chloride 0.5 gm (Anhydrous)

Tartrazine solution:

o Saturated solution of tartrazine in cellosolve (ethylene glycol monoethyl ether),

about 0.3%.

ALDEHYDE FUCHSIN FOR PANCREATIC CELL - HALMI

Procedure: 1. B.S.D.W.

2. Treat with 0.25% acidified potassium permanganate for 5 minutes.

3. Wash in tap water and bleach with 1% oxalic acid.

4. Wash well

5. Rinse in 70% alcohol and stain with aldehyde fuchsin for 5 minutes.

6. Rinse excess stain with 70% alcohol followed by washing with tap water.

7. Counterstain nuclei with Celestine blue-haematoxylin sequence.

8. Rinse with distilled water and stain in Orange G-light green for 1 minutes.

9. Rinse with 0.2% Acetic acid followed by 95% alcohol (no water).


Results: Alpha cell yellow

Beta cell deep purple-violet

D cell and collagen green

Elastic fibre and some mucopolysaccharide purple-violet

Solutions: Halmi’s or Orange G -light green Light green 0.2 g

Orange G 1 g Phosphotungstic acid 0.5 g

Glacial acetic acid 1 ml

Distilled water 100 ml

0.25% acidified potassium permanganate

ALDEHYDE FUCHSIN TECHNIQUE - GORMORI

Procedure: 1. B. S. D. W.

2. Treat with 0.25% Acidified potassium permanganate for 5 minutes.

3. Wash in tap water and bleach with 1% Oxalic acid.

4. Rinse in 70% alcohol and stain with Aldehyde fuchsin for 5 minutes.

5. Rinse quickly with 0.5% acid alcohol.

6. Check microscopically.


Results: HAA positive purplish

Background very pale purplish/colourless

Elastic fibres, sulphated mucin (include mast cells), beta cell of pancreas & pituitary, some

lipofuscins, gastric chief cells hypothalamic neurosecretory cells purple

Solutions: Aldehyde fuchsin Basic fuchsin 0.5 g

70% alcohol 100 ml

Concentrated HCl 1 ml

Paraldehyde 2 ml

Dissolve the dye in the alcohol. Filter and add the acid and paraldehyde. Leave at room

temp. for 72 hours until the stain has assumed a deep purple colour. The stain should be

stored in the refrigerator at 4oC.

BEST'S CARMINE METHOD-BEST, 1906 (BC)

Procedure 1. B. S. D. W.

2. Stain the nuclei with an iron haematoxylin stain. Differentiate and blue.

3. Stain with Carmine solution 5 - 15 minutes. The older the stock carmine used the longer the

staining time.

4. Wash well in either Best's differentiator.

5. Rinse in fresh alcohol.

6. Clear in xylene and mount as desired.

Results: Glycogen deep red Some mucin, fibrin weak red Nuclei blue Solutions:

a) Carmine Stock solution Add 2 g carmine, 1 g potassium carbonate and 5 g potassium

chloride to 60 cm3 distilled water. Boil gently for 5 minutes, using a large flask to avoid

spillage (the heating produces a carimine-metallic ion complex which is the active

staining ingredient). Cool and add 20cm3 concentrated ammonia. Filter and store in a

dark container at 40oC, where it should remain usable for several months.

b) Carmine working solution Stock solution 15 cm3 Concentrated ammonia 12.5 cm3

Methanol 12.5 cm3

c) Best's differentiator Methanol 40 cm3 Ethanol 80 cm3 Distilled water 100 cm3

Remarks: a. Failure to stain is usually due to either using an old deteriorated stock solution, or to

having washed the section in water instead of alcohol or differentiator, immediately

following staining with carmine.

b. An artefact encountered with this technique is stain precipitate on the section. To

minimise this, is it suggested that the following points are noted:

i. The working solution should be filtered before use.

ii. Staining should be carried out in a closed container (e.g. Coplin jar).

iii. The section should not be allowed to dry following staining with carmine, but

should be washed immediately in alcohol or differentiator.

ALIZARIN RED S FOR CALCIUM


2. Flood with Alizarin red solution for 1 minutes.

3. Rinse with 95% alcohol.


Results: Site of calcium deposits orange red Background yellowish

Solution: 2% aqueous solution of Alizarin red S

Adjust pH to 4.1-4.3 with dilute ammonium hydroxide using a glass electrode pH-meter. The

solution should be a deep iodine colour and keeps well at 4oC

BODIAN COPPER PROTARGOL METHOD FOR NERVE FIBRES (BODIAN)


2. Place in copper-Protargol solution at 37 °C for 48 hours.

3. Rinse with distilled water and reduce in hydroquinone solution for 10 min.

4. Wash in distilled water and tone in 0.2% Gold chloride solution for 5 min.

5. Intensify in 2% Oxalic acid until section appears purplish.

6. Rinse in tap water and fix in 2.5%,sodium thiosulphate solution for 3 min.


Results: Nerve fibres, nuclei Greyish black

Background Purplish

Notes: After step (6), section can be counterstained with PAS. No haematoxylin staining of the

nuclei is required.

Solutions: Copper protargol solution

Silver protargol 0.4 g


Copper chips 1 g

Sprinkle the silver protargol powder onto the surface of distilled water and let it dissolve

without stirring. Add copper chips

Hydroquinone solution

Sodium sulphite 1 g

Hydroquinone 0.2 g


CHROMOTROPE-ANILINE BLUE METHOD FOR COLLAGEN AND

MALLORY BODIES (CAB)


2. Stain nuclei with Weigert's iron haematoxylin. Rinse in distilled water.

3. Immerse in 1% phosphomolydbic acid for 1 - 3 min. Rinse well in distilled water.

4. Stain with CAB solution for 8 min. Rinse well in distilled water. Blot.

5. Dehydrate quickly, clear and mount.

Results: Collagen Blue Mallory bodies Blue or sometimes red Giant mitochondria Red

Solution: CAB solution 1.5 g aniline blue is dissolved in 6 ml HCl and 200 ml distilled water with gentle

heat, 6 g Chromotrope 2R is added. The pH should be 1.0.

Remark: As used at Mount Sinai Hospital, New York; modified from Roque (1953) and Churg and

Prado (1956).

BIEBRICH SCARLET-FAST GREEN TECHNIQUE - Guard 1959 (BSFG)


2. Treat with the diluted Harris haematoxylin solution.

3. Drain and transfer direct to the Biebrich scarlet -solution 2 min.

4. Rinse in 50% alcohol.

5. Place in a Coplin jar of fast green solution 12 - 24 hr (overnight at room temperature is

convenient.)

6. Rinse in water, dehydrate, clear and mount as desired.

Results: Nuclear chromatin & sex chromatin body dark green

Cell cytoplasm light green

Solutions: Harris Haematoxylin diluted 1 part with 5 parts distilled water.

Biebrich scarlet solution

Biebrich scarlet ws 1 gm

Phosphotungstic acid 0.3 gm

Acetic acid 0.5 ml 50% ethanol 100 ml

Fast green solution:

Fast green FCF -------------0.5 gm

Phosphomolybdic acid ------0.3 gm

Phosphotungstic acid --------0.3 gm

Glacial acetic acid -----------5 ml

50% ethanol ----------------100 ml

CARBOL FUCHSIN FOR SEX CHROMATIN - BARR BODIES (CF-B)

Procedure 1. Stain in working solution for 1-2 min.

2. Differentiate with 95% ethanol and check microscopically. (½ - 1 min.)

3. D.C.M.

Results: Sex chromatin bodies red (at rim of nuclear membrane)

Solutions: Stock solution Basic fuchsin 3 gm 70% Ethanol 100 ml

Working solution

Stock solution 10 ml 5%

Phenol in D.W. 90 ml

Glacial acetic acid 10 ml

37-40% Formaldehyde solution 10 ml (Stand for at least 24 hrs. Keep for approx. 1

month)

Remark: Medical laboratory technology & clinical pathology. Lynch p. 1280

CRESYL FAST VIOLET FOR NISSL SUBSTANCES

Procedure 1. Section at 8 mm.

2. Stain for 10 min. in cresyl fast violet solution. (It can be heat up to 56oC for rapid result)


4. Differentiate in 95% alcohol until background is relatively clear.

5. D.C.M.

Results: Nissl substances, nucleoli and nuclei violet

Background colourless

Solutions: Cresyl fast violet 0.1%

cresyl fast violet 30 ml

10% glacial acetic acid 5 drops

CRESYL FAST VIOLET - FOR BARR BODY


2. Stain in Cresyl fast violet solution 10 min.

3. Rinse in 0.01N acetic acid.

4. Wash in 0.01N acetic acid.

5. Rinse in absolute ethyl alcohol.

6. Differentiate in absolute ethyl alcohol.

7. Control microscopically until only the sex chromatin body is deeply stained in contrast with

the rest of the nucleus.

8. Clear in xylene, mount in D.P.X.

Results: Sex chromatin body - deep blue

Nucleus - light blue

Notes: A simple method for the demonstration of the sex chromatin body.

Solution Cresyl fast violet 1 gm.

Acetic acid (0.01N), filter 100 ml

DIAZO


2. Treat with the solution for 1 min.

3. Rinse with D.W.

4. Counter stain with Hx.

5. D.C.M.

Result: 1. Argentaffin granules orange red

2. Nuclei blue

3. Background pale yellow

Solution: 4. 1% aqueous fast red salt B 5 cm3

5. Saturated aqueous lithium carbonate 2 cm3

6. Mix and pre cool at 4 °C before use.

CONGO RED FOR AMYLOID (CR)

Procedure 1. Stain nuclei with Harris' Haematoxylin, differentiate and blue.

2. Stain with Congo red solution for 5 minutes.

3. Wash in water and differentiate with alcoholic potassium hydroxide until the background is

clear.

4. Dehydrate, clear and mount..

Results: Amyloid, elastin & eosinophils orange red

Nuclei blue

Background clear

Solutions: 0.5% congo red in 50% alcohol

0.2% potassium hydroxide in 80% alcohol

FEULGEN FOR DNA


2. Treat section with 1 N HCl for 8 minutes at 60 °C.


4. Stain in Schiff's reagent for 45 minutes.

5. Rinse in bisulphite solution.

6. Counterstain with light green for 1 minute.

7. D.C.M.

Results: DNA Magenta Other structure Shades of green

Solution: Light green: 1% light green in 1% acetic acid.

GIEMSA-DIFF Q

Procedure 1. For paraffin section, B.S.D.W. and allow section to dry.

2. For imprints, frozen sections, or smears perform step 3 directly.

3. Dip slide in Solution A 5 times, one sec. each time. Allow excess to drain.

4. Dip slide in Solution B 5 times, one sec. each time. Allow excess to drain.

5. Dip slide in Solution C 5 times, one sec. each time. Allow excess to drain.

6. Wash in tap water.

7. Allow section to dry.

8. Clear & Mount.

Results: Eosinophilic Granules - Red

Nuclei - Blue

Mast Cells - Blue

Cytoplasm - Pink

Bacteria - Blue

FOUCHET FOR BILE PlGMENT


2. Treat with Fouchet reagent 5 minutes.

3. Wash in running tap water 2-3 minutes.

4. Counterstain with Van Geison for 5 minutes.

5. D.C.M.

Results: Bile pigment greenish yellow

Collagen red

Other structures yellow

Solution: Fouchet reagent

25% aqueous trichloroacetic acid 10 ml 1

0% ferric chloride 10 ml

FEULGEN NUCLEAL REACTION FOR DNA - FEULGEN AND ROSSENMCKI

1924

Procedure 1. Bring all sections to water.

2. Rinse sections in HCl at room temperature 1 min

3. Place section in N-HCl at 60oC 8 min

4. Place sections in HCl at R.T. 1 min

5. Transfer sections to Schiff's reagent 45 min

6. Rinse sections in bisulphite solution. 2 min

7. Repeat wash in bisulphite solution. 2 min

8. Repeat wash in bisulphite solution. 2 min

9. Rinse well in distilled water.

10. Counterstain if required in 1% light green.

11. Wash in water.

12. Dehydrate through graded alcohols to xylene and mount.

Results: DNA Red-purple

Cytoplasm Green

Fixation: Not critical but not Bouin.

Reagents: a. N-hydrochloric acid Hydrochloric acid (conc.) 8.5 ml Distilled water 91.5 ml

b. Bisulphite solution 10% potassium metabisulphite 5 ml N-hydrochloric acid 5 ml Distilled

Water 90 ml

GRAM FOR BACTERIA


2. Stain in filtered crystal violet 30 sec.

3. Treat with Gram's iodine one minute.

4. Differentiate with equal parts of acetone and alcohol.

5. Rinse in water for 10-20 minutes or rinse in alcohol to remove the iodine.

6. Counterstain with neutral red.

7. Blot dry and transfer to absolute alcohol.

8. Clear and mount.

Results: Gram positive bacteria - Blue

Gram negative bacteria - Red

Nuclei - Blue

Other structures - Shades of pink

Note: Prolonged treatment with crystal violet will result in staining precipitate though the stain is

filtered. A control section should be stained at the same time.

Solutions: Crystal violet solution: Crystal violet 26 gm 95% alcohol 20 ml Ammonium oxalate 0.8 gm


Gram's iodine: Iodine 1 gm Potassium iodide 2 gm Distilled water 300 ml

Neutral red: 1% aqueous solution

GRIMELIUS FOR ARGYROPHIL CELL

Procedure 1. Place the slides in the freshly prepared silver solution, for 5 hours at 60 °C.

2. The silver solution was drained off from the slides (not the section) and these were then

immersed in a freshly prepared reducing solution for 1 minute at 60 °C.

3. D.C.M.

Results: Argyrophil Granules : Brown / black

Note: 1. In case where the argyrophil islet cells appeared weak the staining reaction could be

improved by double impregnation. After staining step 2, the slides were immersed in an

aqueous solution of 5% sodium thiosulphate for 2 minutes. After rinsing in distilled water for

about 5 minutes the slides were placed in a further freshly prepared silver solution as in step

1 for 10 minutes at room temp. and then in a freshly prepared reducing solution as in step 2

for 1 minute at 40-45oC. 2. For step 1, the slides can be incubated at 37oC for overnight

Solutions: Silver solution: 0.1 M acetic acid-sodium acetate buffer pH 5.6 3.3 ml (0.2 M Acetic acid 19

ml, 0.2 M sodium acetate buffer 181 ml) Distilled water 30 ml Freshly prepared 10% silver

nitrate 0.1 ml

Reducing solution: Hydroquinone 0.4 gm Sodium sulphite 2 gm Distilled water 40 ml

GROCOTT METHENAMINE SILVER NITRATE METHOD WITH

MICROWAVE TECHNIQUE


2. Place section in 5% Chromic acid solution at 60 °C for 120 seconds (time at temp).

3. Wash section in tap water.

4. Treat section with 1% Sodium metabisulfite solution for 60 seconds.

5. Rinse section in tap water followed by several changes of distilled water.

6. Place section in Methenamine silver working solution and heat until the solution starts to

boil (time at temp at 100oC for 2 seconds).

7. Leave section in hot working solution until proper staining intensity is achieved.

8. Rinse section in distilled water. * If additional incubation is required, place section back in

hot working solution.

9. Tone section in 0.2% Gold chloride solution for 5 minutes.

10. Rinse section in distilled water.

11. Treat section with 2.5% Sodium thiosulphate solution for 2 minutes.

12. Wash section in tap water.

13. Counterstain section with 1% light green solution.


Results: Fungi Sharply outlined in black

Nuclei Grey

Background Pale green

Solutions: 3% Methenamine solution

o Methenamine 3 g

o Distilled water 100 ml

5% Aqueous silver nitrate solution

o Silver nitrate 5 g


GRIMELIUS METHOD WITH MICROWAVE TECHNIQUE

Procedure 1. Prepare silver nitrate and reducing solution.

2. B.S.D.W.

3. Incubate section in silver nitrate solution and heat to 100oC for 5 seconds.

4. Leave section in (3) for 1 minute.

5. Preheat the reducing solution at 100oC for 5 seconds.

6. Rinse the section with distilled water.

7. Reduce section in (5) until yellowish brown colour macroscopically.

8. Rinse section with distilled water.


Results: Argyrophil granules brown / black

Solutions: Silver nitrate solution

o silver nitrate 0.5 g


o Acetate buffer pH 5.6 5 ml

Reducing solution

o Sodium sulphite 2 g

o Hydroquinone 0.4 g

o Distilled Water 40 ml

GROCOTT'S METHENAMINE SILVER NITRATE METHOD FOR FUNGI


2. Oxidize in 5% Chromic acid for half an hour.

3. Wash in running tap water.

4. Rinse for 2 minutes in 1% Sodium metabisphite to remove residual chromic acid.

5. Wash in tap water, rinse thoroughly in distilled water.

6. Treat with Methenamine silver nitrate solution at 58oC for half an hour. (check

microscopically)

7. Rinse thoroughly with distilled water.

8. Tone in 0.1% Gold chloride for 5 minutes.


10. Treat with 2.5% Sodium thiosulphate for 2 minutes.

11. Wash thoroughly with distilled water.

12. Light green counterstain.


Results: Fungi Sharply outlined in black Nuclei Grey Background Pale green

Solutions: 5% Chromic acid Chromic acid 5 g Distilled water 100 ml Methenamine silver nitrate solution

3% Methenamine(Hexamine) 10 ml 5% Silver nitrate 0.5 ml 5% Borax(std.borate) 0.8 ml


Freshly prepared, filter before use.

5% Borax solution Borax 5 g Distilled water 100 ml

Methenamine silver stock solution 3% Methenamine solution 400 ml 5% Aqueous silver

nitrate solution 20 ml

Mix slowly, kept in brown bottle and store at 4oC.

Methenamine silver working solution Stock solution 20 ml 5% Borax solution 1.6 ml Distilled

water 20 ml

GOMORI'S TRICHROME - RAPID, ONE STEP METHOD

Procedure 1. Fix air dry frozen section in buffered formalin for 15 minutes.

2. Stain nuclei with an iron haematoxylin.

3. Differentiate and, blue as per the standard technique.

4. Wash well in tap water, then in distilled water.

5. Stain in Gomori solution for 5 to 20 minutes.

6. Rinse well in 0.2% glacial acetic acid solution.

7. Blot dry, dehydrate, clear and mount.

Results: Nuclei grey-blue Collagen green Muscle, cytoplasm, red blood cells, fibrin red Higher pH

Nemaline rods red Background blue-green

Solution: Gomori solution Chromotrope 2R 0.6 g Fast green FCF 0.3 g Phosphotungstic acid 0.6 g

Glacial acetic acid 1 ml Distilled water 100 ml

Note: For demonstrating nemaline rods, adjust pH of the solution to 3.4 using 1 M NaOH

GORDON AND SWEET'S FOR RETICULIN


2. Treat with 0.25% potassium permanganate solution 5 minutes.

3. Rinse in tap water.

4. Bleach in 1% oxalic acid solution 5 minutes.


6. Treat with 2.5% iron alum for at least 15 minutes.

7. Rinse well with distilled water.

8. Treat with silver bath 20 seconds or above (determined by the age of the silver bath).


10. Reduce with 10% formalin solution for 1 - 2 minutes.

11. Rinse in tap water.

12. Fix in sodium thiosulphate for 1 - 2 minutes.

13. D.C.M.

Results: Reticulin Black Collagen Golden brown Nuclei Black

Reagents: Silver bath 10% silver nitrate 5 ml. Titrate with ammonia dropwisely until solution is just

about to clear. Add in 5 ml 3% sodium hydroxide and titrate with ammonia until solution is

just about to clear. Make up to 50 ml with distilled water. Filter before use.

Notes: Rinse all the apparatus thoroughly in distilled water before use.

HAEMATOXYLIN-BASIC FUCHSIN-PICRIC ACID FOR MYOCARDIAL

INFARCTION


2. Stain nuclei with Harris' haematoxylin for seconds.

3. Wash in water, blue.


5. Stain in 0.1% Basic Fuchsin for 3 minutes.

6. Rinse in distilled water, blot.

7. Rinse in acetone.

8. Differentiate in 0.1% Picric acid in acetone for 5 seconds.

9. Stop differentiation with rinse in acetone.

10. Clear in xylene and mount.

Results: Nuclei Blue Ischemic Muscle, RBCs Red Normal Muscle Yellow

Solutions: a) Harris' Haematoxylin.

b) 0.1% aqueous Basic Fuchsin (C.I. 42510).

c) Acetone.

d) 0.1% Picric Acid in Acetone.

GOMORI'S MODIFIED TRICHROME - ENGEL AND CUNNINGHAM

Procedure 1. Use air-dried, cryostat-cut sections (10mm).

2. Stain in Harris's haematoxylin for 3 min.

3. Rinse briefly in distilled water.

4. Decolorize in 0.5% acid alcohol.

5. Blue in Scott's tap water.


7. Stain in Gomori’s stain for 10 min. Check microscopically.

8. Rinse BRIEFLY in 0.2% acetic acid.

9. Dehydrate quickly, clear in xylene and mount with DPX.

Results: Intermyofibrillar material red Myofilaments green Nuclei reddish blue Connective tissue

green

Solutions: Gomeril’s stain Chromotrope 2R 0.3 gm Fast Green FCF 0.15 gm Phosphotungstic acid 0.3 gm

Glacial Acetic acid 0.5 ml

Add distilled water to make 50 ml. Adjust to pH 3.4 with 1N NaOH

Note: This stain must he freshly prepared and sections are stained after cutting.

Remarks: 1. Particularly helpful in identifying nemaline rods.

2. Lillie Page 701

HEIDENHAIN'S IRON HAEMATOXYLIN (H.I.H.)

Procedure 1. Bring section to water.

2. Place in iron alum mordant for 30 min.


4. Place section in haematoxylin solution for 30 min.

5. Rinse well in tap water.

6. Differentiate in 2.5% iron alum solution until desired staining is achieved.

7. Wash in tap water for 10 min. to remove the iron alum.

8. Counterstain with sat. tartrazine in absolute cellosolve for 30 sec.

9. Rinse with 95% alcohol. D.C.M.

Results: Nuclei, muscle striations, RBC, keratin - Bluish Black Background - Yellowish

Solution: Iron Alum Mordant 5% aqueous ferric ammonium sulphate (iron alum)

Heidenhain's Haematoxylin Solution Haematoxylin 0.5 gm Absolute alcohol 10 ml Distilled

water 90 ml

Iron Alum Differentiater 2.5% aqueous iron alum solution.

HAEMATOXYLIN EOSIN (H&E)

Procedure 1. Section down to water.

2. Stain in Haematoxylin solution for 5 minutes.

3. Rinse well in tap water.

4. Differentiate in 0.5% acid alcohol.

5. Blue in 1% scott's solution for 1 minute.

6. Rinse briefly.

7. Stain in Eosin solution for 1 minute.

8. Rinse briefly in D.W.

9. D.C.M.

Results: Nuclei Blue Background Pinkish to orange red

Reagents: HARRIS HAEMATOXYLIN:

Haematoxylin 8 gm Absolute alcoholol 80 ml Ammonium alum or potassium alum 160 gm

Distilled water 1600 ml Glacial Acetic Acid 32 ml Mercuric oxide 2 gm

SCOTT'S TAP WATER:

Sodium Bicarbonate 7 gm Magnesium Sulphate 40 gm Dissolve in 2 litres of distilled water

Thymol is added to prevent mould formation

ACID ALCOHOL:

Conc. HCl 5 ml 70% Alcohol 995 ml

EOSIN SOLUTION:

10% Eosin 60 ml 1% Erythrosin 40 ml Distilled water 500 ml

LAQUEUR - ALCOHOLIC HYALINE (Laqueur)

Procedure 1. Stain in alum Hx for 5 minutes.

2. Stain in acid fuchsin solution and heat with a small flame to fuming. Let stand 5 minutes.

3. Wash, differentiate in a mixture of 7 parts 20% alcohol and 1 part sat. Alcoholic picric acid

until only hyaline and red corpuscles remain red, and collagen is faint grey or unstained.

5. Wash thoroughly.

6. Mordant 30 minutes in 1% phosphomolybdic acid.

7. Transfer directly to 1% light green SF in 1% acetic acid until collagen stains green.

8. D.C.M.

Results: Mallory's alcoholic hyaline Brilliant red Cytoplasm Pale brown Bile pigment, collagen Green

Haemosiderin and hemofuscin Unstained/yellowish brown

Solution: Acid fuchsin solution : Shake 2 ml of aniline with 100 ml D.W. Filter. Dissolve 20 g acid

fuchsin in aniline water prepared.

IRON HAEMATOXYLIN STAIN FOR AMOEBAE CHROMOSOME

STRUCTURE - CRAIG AND FAUST, CLINICAL PARASITOLOGY, P.864 (IH)

Procedure 1. Fix smears in Schaudinn's solution at 60oC for 2 minutes.

2. Remove Hg by 70% alcohol with iodine.

3. wash in water.

4. Mordant in 2% iron alum at 40oC for 2 minutes.

5. Wash in water for 3 minutes.

6. Stain in 0.5% aqueous Hx. 2 minutes.

7. Differentiate in iron alum until chromosomes are prominent.

8. Wash in water.

9. D.C.M.

Solution: Schaudinnls solutions Solution A:

Sat-aq. HgCl2 600 ml 95% ethyl alcohol 300 ml before use add 5 ml acetic acid to every 100

ml solution A

SOUTHGATE’S MUCICARMINE (MC)

Procedure 1. Stain nuclei with Harris Hx for 5 minutes, differentiate and blue.

2. Treat with Mucicarmine solution for 20 minutes.

3. Rinse in water.


Results: Epithelial mucin red Cryptococci red Nuclei blue

Soultion: Mucincarmine (stock) Carmine 1 g Aluminium hydroxide 1 g 50% alcohol 100 ml Aluminium

chloride 0.5 g (0.9 g if A1C13.6H20) is used.

After grading Carmine into paste with 100 ml 50% alcohol, then add 1 g Al (OH)3 and 0. 9 g

A1C13. 6H20. Mix and boil gently for 2.5 minutes, cool and filter, and store in the

refrigerator. When use, the stain is diluted 1:10 (The optimal dilution factor has to be tried

and varies from batch to batch.)

MANN’S METHYL BLUE - EOSIN FOR MINUTE STRUCTURE AND

INTESTINAL FLAGELLATE FLAGELLATE OF AMOEBAE( NW)

Procedure: 1. 1 Fix smears in Schaudinn's fluid 60oC 2 minutes.

2. Remove Hg pigment by 70% alcohol with iodine.

3. Stain for 4 - 12 hours in staining solution, as determined by trial.

4. Wash in distilled water thoroughly.

5. 5 Differentiate in 70% alcohol containing a little OG (a few drops to 100 ml.)

6. Wash in water.

7. D.C.M.

Solutions: 1% Methyl Blue 35 ml 1% Eosin 45 ml Distilled water 100 ml

Notes : Craig and Faustm, Clinical Parasitology, p.864

MARTIUS YELLOW - SCARLET RED, SOLUBLE BLUE FOR FIBRIN (MSB)

Procedures: 1. Stain nuclei with celestine blue - Haemalum sequence, 3 minutes each, differentiate and

blue.

2. Rinse in 95% alcohol and stain with 0.5% martius yellow for at least 2 minutes. Rinse in 95%

alcohol. Cheek that only RBC is stained yellow.

3. Stain in 1% brilliant crystal scarlet 1 minute, check microscopically.

4. Treat with 1% phosphotungstic acid for a short while and check that only muscle and fibrin

are stained red.

5. Counter stain with 0.5% soluble blue for 1 minute. (Be careful of overstain).

6. D.C.M.

Results: Fibrin and Muscle Red RBC Yellow. Nuclei Greyish blue Collagen Blue

Solutions: 0.5 % Martius yellow :

Martius yellow 0.5 gm Phosphotungstic acid 2 gm 95 % Alcohol 100 ml

1 % Brilliant crystal.scarlet : Brilliant crystal scarlet 6R 1 gm Glacial acetic acid 2.5 ml

Distilled water 97.5 ml

0.5 % Soluble blue

MASSON FONTANA (MF)

Procedures: 1. 1 B. S. D. W.

2. Rinse section thoroughly with distilled water.

3. Stain overnight in Fontana silver solution at dark in a closed jar.


5. Fix in 0.5% sodium thiosulphate for 2 minutes.

6. Wash in running tap water for 2 minutes.



Results: Melanin black Argentaffin granule black nuclei red

Solutions: Fontana silver solution

Add strong ammonia to 20 ml of 10% aqueous silver nitrate, until the precipitate dissolves.

Add 20 ml of distilled water and filter before use.

TAEZER-UNNA ORCEIN METHOD (ORCEIN)


2. Bleach with acidified KmnO4 for 5 minutes, then oxalic acid 1 min.

3. Take section to 70% alcohol.

4. Place in a closed jar of the orcein stain for 2 hours for HAA at room temp.

5. Differentiate with 1% acid alcohol.

6. Stain briefly in Hx for 10 sec.

7. Wash well in tap water for blueing.

8. D.C.M.

Results: Elastic fibers dark brown Nuclei blue HAA purple brown

Solutions: a) Orcein stain Orcein (synthetic) 1 g 80% alcohol 100 ml ( adjust to pH 1 - 2 with HCl)

conc. HCl 1 ml

b) 1% acid-alcohol

c) Methylene blue or alum haematoxylin

d) Acidified KMn04 : 0.25% KMn04 10 ml conc. H2S04 12 ml

MASSON TRICHR0ME

Procedures 1. Weigert’s haematoxylin 5 minutes (or Celestine Blue - Hx sequence, 5 min : min).

2. Cytoplasmic stain 5 minutes (optimum)

3. Differentiate with 1% phosphomolybdic acid, check under microscope.

4. Counterstain with 1% light green. Be careful of over staining.

5. . Rinse with 1% acetic acid and then 95% alcohol.


Results: Muscle, keratin, RBC red Nuclei bluish black Collagen, basement membranes, reticulin,

elastic fibres green

Solutions: Celestine Blue Celestine blue 0.5 g Ferric ammonium sulphate 5 g Glycerin 14 ml Distilled

water 100 ml

Dissolve the iron alum in the water without heat. Add the celestine blue and boil for 3

minutes. Filter when cool and add the glycerin. The stain can keep for at least 6 months.

Cytoplasmic stain 1% ponceau 2R in 1% acetic acid 2 parts 3% acid fuchsin in 1% acetic acid 1 parts

Fibre stain 2% Light green in 1% acetic acid Commercial Gomeri’s trichrome stain may also be used,

stain for 2-5 minutes, rinse in 1% acetic acid. result is the same for MT.

PALMGREN’S METHOD FOR NERVE FIBRES (PALMGREN)

Procedure 1. Cut paraffin section at 7 microns and celloidinize section.

2. Treat with acid bath for 5 min.

3. Rinse with 3 changes of distilled water for 5 min.

4. Stain with silver bath for 30 min. at room temp.

5. Drain off excess solution and shake continuously in reducing bath at 45oC for 1 min.

6. Rinse with 50% alcohol for 10 seconds.

7. Wash with distilled water, 3 changes in 5 min.

8. Intensify in 2% oxalic acid until section appears dark brown.

9. Treat with 2.5% sodium thiosulphate for 3 min.

10. D.C.M.

Results: Nerve fibres and neurofibrils Black Background Yellowish brown

Solutions : Acid Bath: 40% Formaldehyde 25 ml Distilled water 75 ml 1% Nitric acid 0.2 ml

Reducing Bath: Pyrogallol 1 gm Distilled water 45 ml Absolute alcohol 55 ml 1% Nitric Acid 0.2 ml

Silver Bath: 15% silver nitrate

OIL RED O METHOD FOR LIPIDS (ORO)

Procedure: 1. Cut frozen section at 6 mm.

2. Air dry at room temp. for 1 hour and rinse with 60% triethyl phosphate and fix in formalin

for 10 minutes.

3. Filter working solution onto section and leave in moist chamber for 15 minutes

(Alternatively, it can be stained in a coplin jar which contains 6 part dye and 4 part DW for 15

- 30 minutes).

4. Pour off excess stain and differentiate in 60% triethyl phosphate solution for a few seconds.

5. Bring section to water.

6. Counterstain nuclei with Harris Haematoxylin for 15 second.

7. Blue nuclei. Do not differentiate in acid alcohol.

8. Mount in aqueous mountant.

Results: Neutral lipids orange red

Nuclei blue

Note: Formalin fixed tissue block can be placed in gum sucrose solution overnight before the

frozen section. Use the free floating method or attach the section to albuminized slides.

Gum sucrose solution :

Gum acacia (gum arahic) 2 gm Sucrose 60 gm Distilled water 200 ml

Completely dissolve the gum acacia in water with stirring. Add sucrose and dissolve. Store at

4oC.

Reagents: Stock solution : sat. oil red 0 in triethyl phosphate.

PASM FOR BASEMENT MEMBRANE(PASM)


2. Oxidize in 1% Periodic acid for 15 minutes.

3. Oxidize in 5% Chromic acid for 15 minutes.

4. Wash in running water.

5. Rinse in 1% Sodium bisulphite to remove residual chromic acid.

6. Wash in tap water, rinse thoroughly in distilled water.

7. Treat with Methenamine silver nitrate solution at 58oC for half an hour. (check

microscopically)


9. Tone in 0.1% Gold chloride for 5 minutes.


11. Treat with 2.5% Sodium thiosulphate for 2 minutes.


13. Light green counterstain


Solutions: Methenamine silver solution

3% Methenamine 15 ml 5% Silver nitrate 0.75 ml 5% Borax 1.2 ml Freshly prepared, filter

before use.

Results: Basement membrane black Background green

PERIODIC ACID-SCHIFF REACTION (PAS)


2. Treat with 1% Periodic acid for 5 minutes.


4. Treat with Schiff reagent for 5 minutes.


6. Put in running water for at least 5 minutes.

7. Counterstain with Harris' haematoxylin for 2 minutes.

8. D. C. M.

Results: PAS positive substances magenta negative after digestion glycogen nuclei blue

Solutions: Basic fuchsin 1 g 0.15 M HCl 100 ml (1.275 ml of conc. HCl in 100 ml distilled water)

Sodium metabisulphite 2 g Dissolve Basic fuchsin in 0.15 M HCl with stirring. Add sodium

metabisulphite and stir for 8 hours until the solution appears straw colour. Add activated

charcoal (2 spoonful) to absorbs the colour. Filter and store aliquots at 4oC

Note: Treat section with amylase for 15 minutes at room temperature before step (2) for PAS with

digestion (PASD) method.

PHLOXINE TARTRAZINE (PT)


2. Stain nuclei with Harris Hx.

3. Stain in phloxine solution for 10 min.


5. Differentiate in tartrazine solution, controlling microscopically until a yellow background

with red keratin and RBC.

6. Rinse in 95% alcohol (never rinse in water).

7. D.C.M.

Results: Keratin, RBC, Paneth cell granules, fibrin, certain inclusion bodies red Nuclei greyish blue

Other structures yellow

Solutions: Phloxine solution: Phloxine 0.5 g Calcium chloride (anhydrous) 0.5 g Distilled water 100 ml

Tartrazine solution :

Saturated solution of tartrazine in cellosolve, about 0.3%.

PERLS METHOD FOR HAEMOSIDERIN (PERLS)


2. Rinse thoroughly in distilled water.

3. Stain in Perls' reagent for 25 min.


5. Counterstain with 1% Neutral red for 2 min.

6. Wash in 95% alcohol.


Results: Haemosiderin, Ferric ions Blue Nuclei Red

Solutions: Perls’ reagent

Equal parts of 2% potassium ferrocyanide and 2% hydrochloric acid 1% Neutral red

Neutral red 1 g Distilled water 100 ml

RUBEANIC ACID FOR COPPER (RA)


2. Stain in rubeanic acid solution overnight at 37oC.

3. Place section in 70% alcohol for 15 minutes.

4. Counterstain with Van Gieson solution for 3 minutes.


6. D.C.M.

Results: Copper greenish black granules Collagen red Background yellowish

Solutions: Rubeanic acid solution

0.1% Rubeanic Acid in Absolute Alcohol 2.5 ml 10% Sodium Acetate 50 ml

In QAP, Rubeanic acid solution 0.1% rubeanic acid in 70% alcohol 1 parts 10% sodium

acetate 20 parts

MALLORY PTAH METHOD FOR MUSCLE STRIATIONS (PTAH)


2. Treat section with 0.25% potassium permanganate minutes.

3. Bleach with 1% oxalic acid until section is clear.

4. Mordant with 2.5% iron alum for 1 hour.

5. Stain in PTAH solution overnight.

6. Rinse briefly with 95% alcohol.

7. D.C.M.

Results: Nuclei, Fibrin, Muscle Striations, Myofibrils, RBC, Astroglia Blue

Collagen, Bone and Cartilagenous Matrix Orange red

Solutions: PTAH solution

Haematein 1 gm Phosphotungstic Acid 10 gm Distilled water 1000 ml

Dissolve the haematein in one half of the water with a little of heat. The phosphotungstic

acid in the other half of water and then mix. Store in a tightly stoppered bottle and allow the

solution to ripen.

SCHMORL'S FERRICYANIDE (SF)

Procedures: 1. Wash section well in distilled water.

2. Stain in ferric ferricyanide solution for 10 minutes.

3. Rinse in 1% acetic acid.

4. Rinse in water.

5. Counterstain in Van Gieson solution for 1 minute.

6. Rinse with 95% alcohol.

7. D.C.M.

Results: Melanin, intestinal argentaffin lipofuscin, chromatin blue Cytoplasm, RBC, muscles yellow

Collagen red

Regents: Ferric ferricyanide solution :

1% aq. ferric chloride 30 ml 1% potassium ferricyanide 4 ml distilled water 6 ml

The solution has to be freshly prepared.

SOLOCHROME CYANIDE FOR MYELIN (SC-M)


2. Stain in solochrome cyanide solution for 10-20 minutes at R.T.

3. Wash in running water.

4. Differentiate in 10% iron alum until nuclei are scarely visible.


6. Counterstain if desired.


Results: Myelin sheaths bright blue RBC blue Nuclei pale blue to almost unstain

Solutions: Solchrome cyanide RS 0.2 gm D.W. 96 ml 10% iron alum 4 ml conc. H2S04 0.5 ml

TOLUIDINE BLUE - FOR. DEMONSTRATION OF METACHROMASIA IN

FROZEN SECTIONS (TB-F)

Procedure 1. Fix sections in an acetone: tetrahydrofuran mixture (1:1 by volume) at room temperature for

20 min.


3. Stain with toluidine blue solution for 2 min.


5. Clear in xylene and mount.

Results: Nnuclei blue Mucopolysaccharide red to purple

Solutions: Toluidine blue solution 0.5% toluidine blue in 25% acetone:

1% toluidine blue 5 ml acetone 2.5 ml D.W. 2.5 ml

VERHOEFF (VERHOEFF)


2. Stain in Verhoeff’s reagent for at least 1 hour (a longer incubation time will give a better

result).

3. Rinse in water.

4. Differentiate in 2% ferric chloride, check under microscope until only elastic fibres and nuclei

are black.

5. Rinse in water or rinse in alcohol to remove.iodine.

6. Counterstain with Van Gieson for 1 minute.

7. Blot dry, rinse in 95% alcohol. D.C.M.

Results: Elastic fibres Black Collagen Red Nuclei Grey Other structures Yellow

Solutions: Verhoeff reagent:

5% alcoholic Hx 2 ml 10% ferric chloride 8 ml Iodine 8 ml

Verhoeff iodine :

Iodine 2 g Potassium Iodide 4 g Distilled Water 100 ml

5% alcoholic Hx:

Haematoxylin 5 g Absolute alcohol 100 ml

VICTORIA BLUE (VICTORIA)


2. Oxidize with 0.25% potassium permanganate for 10 minutes.

3. Reduce in 5% oxalic acid for 3 minutes.


5. Wash briefly in 95% alcohol and transfer to the Victoria Blue staining solution for 18 to 24

hours (overnight).

6. Wash in 95% alcohol until excess stain is removed.

7. Counterstain with VG for 1 minutes - Blot dry.

8. Clear and mount.

Results: Elastic fibres Blue-black Collagen Red Other tissues Yellow

Solutions: Victoria Blue staining solution

Victoria blue 4R (C.I. 42563) 1 g Ethyl violet (C.I. 42600) 1 g Dissolve in 200 ml of boiling

distilled water. Add, in the following order :

Resorcin 4 g Dextrin 0.5 g 30% ferric chloride (freshly prepared) 25 ml

Boil for 3 minutes, cool rapidly and filter. Wash precipitate well with distilled water. Transfer

precipitate plus filter paper to the original beaker. Add 100 ml 95% alcohol to precipitate

and boil gently in water bath for 15 minutes. Cool, filter and make up to 350 ml with 95%

alcohol. To this solution add 10 g phenol and 4 ml concentrated hydrochloric acid.

VAN GIESON (VG)


2. Celestine blue 5 minutes, rinse in water, Harris Hx 5 minutes, rinse in water.

3. Differentiate and blue.

4. Stain in Van Gieson - stain 2-3 minutes.

5. Blot dry, rinse in 95% alcohol, D.C.M.

Results: Collagen red Nuclei grey Other structures yellow

Notes: Water removes the red colour, alcohol removes the yellow colour, so that dehydration

should be rapid. 2. Step 1 can be replaced by :

1) Weigert's hx Solu A : Solu B = 1:1 5 minutes

2) briefly differentiated by Solu B.

3) Rinse with water.

4) Van gieson stain.

Solutions: Van Gieson stain : 1% aqueous acid fuchsin 13 ml sat. aqueous picric acid 100 ml

WADE-FITE FOR M. LEPRAE (WF)

Procedure 1. Dewax section in a mixture of equal parts of liquid petrolatum and rectified turpentine

(Terpineol or xylene), blot until almost dry.

2. Wash in tap water, 5 minutes.

3. Stain in carbol-fuchsin solution for 30 minutes.

4. Wash in tap water, blot dry.

5. Decolorize in 3% sulphuric acid.

6. Wash in tap water.

7. Counterstain 0.1% methylene blue.

8. Wash in water, blot dry, xylene, mount.

Results: Positive - Magenta colour

Note: Bleach can be done for clearer background. (potassium permanganate and oxalic acid bleach

after step 6).

VON KOSSA (VK)


2. Rinse section well with distilled water.

3. Treat with 5% silver nitrate solution under strong light for 60 minutes.


5. Fix in 2.5% sodium thiosulphate for 1 minute.


7. D.C.M.

Results: Insoluble calcium salts (also urates) brown to black Other structures shades of pink

Notes: A negative control can be used by treating a parallel section with 2% HCl for 30 min before

staining in step (2).

ZIEHL-NEELSEN (ZN)


2. Flood section with carbol fuchsin stain 10 minutes.

3. Heat slide until steaming, stain for 10 minutes.


5. Decolourize with 1% acid alcohol until section is faintly pink.

6. Rinse thoroughly with tap water.

7. Bleach with 0.25% potassium permanganate, then 1% oxalic acid.


9. Flood slide with 0.25% methylene blue for ½ minute.


Results: Acid fast bacilli red Nuclei blue

Note: 1) A control should always be stained at the same time.

2) Methylene blue can be easily decolourised in alcohol water mixture.

3) Never place stained section in xylene for too long.

4) Long ZN : 3 hours 60oC

Reagents: Carbol fuchsin stain : Basic fuchsin 10 g Alcohol 100 ml 5% aqueous phenol 1000 m1

5% Aqueous phenol: phenol 50 g Distilled water 1000 ml

WARTHIN-STARRY


2. Wash sections well in buffer solution (a).

3. Stain sections in silver solution (b) for 1 hour at 60oC.

4. Develop sections in developer solution (c) for about 1 minute at 60oC until sections appear

yellowish brown.

5. Rinse in tap water at 60oC.

6. D.C.M.

Results: Spirochaetes black Blackground : pale yellow to brown.

Solutions: a) Buffer solution Sodium acetate 8.2 g Acetic acid 12.5 g Distilled water 1000 ml

b) Silver solution Silver nitrate 0.2 g Buffer solution 20 ml

c) Developer solution Solution 1 Hydroquinone 0.06 g Buffer 2.0 ml

Solution 2 Gelatine 1.5 g D.W. 30 ml

Solution 3 Ag nitrate 0.12 g D.W. 6 ml

Notes: Solution 1 mix with solution 2 first, then add solution 3 immediately before use.

Celloidinization

When section is brought to absolute alcohol after deparaffinization, it is placed in 1%

celloidin in equal parts of ether and absolute alcohol for a few seconds. It is then withdrawn

slowly and steadily. The section is then air dried before being transferred to 70% alcohol to

complete the rest of the staining procedures.

LILLIE’S MAYER HAEMATOXYLIN (LILLIE)

Procedure: Dissolve alum in boiling and boil distilled water. Add haematoxylin and boil for 2 minutes.

Add sodium iodate and stir. Colour change to purplish. Cool and add glycerin and glacial

acetic acid. Filter before use.

Reagents Haematoxylin 10 gm Aluminum ammonium sulphate 100 ml Sodium iodate 0.6 gm Glycerin

600 gm Distilled water 1400 ml Glacial acetic acid 40 ml

WEIGERT’S IRON HAEMATOXYLIN

Solution A Haematoxylin 1 g

Ethyl alcohol 100 ml

Solution B 30% aq. Ferric chloride 4 ml

Conc. hydrochloric acid 1 ml


Mix equal volumes of A and B prior to use.

MAYER’S EGG ALBUMIN - GLYCEROL

Solutions: Egg white 50 ml

Glycerol 50 ml


Thymol 100 mg

Mix thoroughly, filter through several layers of gauze into bottle. Store at 4oC.

WHOLE PREPARATION OF WORKS (WORM)

Procedure: 1. Straighten the worm by shaking vigorously for a few seconds in normal saline.

2. Fix in 70% alcohol with 3% acetic acid.

3. Stain for 12 hours or longer in Kirkpatricks carmalum.

4. Differentiate in acid alcohol (may take several hours).

5. Dehydrate in ascending alcohol (50, 70, 95%) leaving for at least half an hour in each grade.

6. Complete dehydration in absolute alcohol, to overnight.

7. Clear in beechwood creosote (cedarwood oil). 8. Mount in balsam. Harden the mount on a

hot plate.

Solutions: Kirkpatricks carmalum: Carmine 5 g Glacial acetic acid 5 ml Potassium alum 5 g D.W. 200 ml

Notes: Grind the carmine in a morter, add acetic acid and 20 ml of water, allow to soak for 20

minutes, then boil gently for 1 hour. Dissolve alum in 100 ml of water and add it to the

cooled carmine solution. Boil gently for another hour. Cool and filter. Add 0.2 g salicylic acid

to inhibit growth of moulds. Stain keeps for at least 1 year.

ORCEIN FOR HEPATITIS B SURFACE ANTIGEN


2. Incubate in 1% potassium permanganate for 5 minutes.

3. Wash in water.

4. Decolourise in 2.5% oxalic acid.

5. Stain in orcein solution for 2-4 h at room temperature.

6. Rinse in 70 % ethanol.

7. Rinse in water, then differentiate in 3% acid alcohol.


Results: HBSAg, copper-associated protein Brown and elastic fibres The method is suitable for

routine demonstration of HBsAg, though probably less sensitive than most

irnmunohistochemical methods.

Solutions: Acidified potassium permanganate

To 95 ml of 0.5% potassium permanganate add 5 ml of 3% sulphuric acid.

Orcein solution

Orcein 1.0 g 70% Ethanol 100 ml Concentrated Hcl 2.0 ml

The pH of the stain should be 1.0 to 2.0. The solution keeps for 1-4 weeks.

Different orcein vary in staining properties (Kirkpatrick 1982). Natural orcein from British

Drug Houses is suitable. Other orcein may require twice the concentration and twice the

amount of HCL (Poppe.personal communication).

GIEMSA STAIN (1 : 1000)

Solutions: 1. Giemsa Stock solution :

a. Giemsa Stain powder 4 g

b. Glycerol 250 ml

c. Methanol (pure) 250 ml

The powder is dissolved in the glycerol at 60oC with regular shaking. After the methanol was

added, the mixture well shaken, and then allowed to stand for 7 days. Filter before use.

2. Working Giemsa stain :

a. Giemsa stock solution 4 ml

b. Buffered distilled water (pH 6.8) 96 ml

Procedure: 1. Sections to water.

2. Rinse in buffered distilled water (pH 6.8).

3. Stain in working Giemsa stain overnight.


5. Differentiate in 0.5% acetic acid until the section is pink.

6. Wash in tap water,rinsed in distilled water

7. Blot until almost dry.

8. Dehydrate very rapidly through alcohols, clear in xylene and mount in DPX.

Results: Eosinophil granules, R.B.C. orange-red Nuclei blue Background pink

Notes: To achieve a good colour balance, it is necessary to initially overstain with the Giemsa stain,

and then slightly over-differentiate in weak acetic acid until there is an overall pink cast to

the cells. -This is come to offset the loss of eosinophilia and gain in basophilia which result

upon eventual alcohol dehydration.

METHYL GREEN PYRONIN

Solutions: 2% Methyl Green (chloroform washed) 9 ml

2% Pyronin Y 4 ml

Acetate buffer pH 4.823 ml

Glycerol 14 ml

Mix well before use.

Procedure: 1. section to distilled water

2. stained with the solution for 30 mins.

3. washed in distilled water

4. differentiate with ethanol

5. cleared and then mounted

Results: RNA (plasma cell cytoplasm, nucleolus) Red DNA (nuclei) Green

Notes: Plasma cells are found in lymphoid tissue, spleen, bone marrow. The plasma cells have an

ovoid cytoplasm and eccentric nucleus with prominent radial chromatin. They are derived

from B lymphocytes and secrete immunoglobulins

This method demonstrated both nuclei acids - methyl green is a basic dye and combination

with the phosphoric groups of DNA and is generally held to be specific -pyronin staining of

RNA is not specific, as it stains some mucous cells and depolymerized DNA - the pH of the

staining solution is critical as well-as conc. of the 2 dyes

The differentiation is important Washing in water after staining is avoided Tissues that have

been decalcified in acids are unsuitable Loyez’s (1910)

HAEMATOXYLIN METHOD FOR MYELIN (AFTER ANDERSON, 1929)

Preparation of staining solutions: a. 4% aqueous iron alum

b. Loyez haematoxylin 10% alcohol haematoxylin 10 ml distilled water 90 ml saturated aqueous

lithium carbonate 4 ml

c. Final differentiator Sodium tetraborate (Borax) 2 g Potassium ferricyanide 2.5 g Distilled

water 200 ml

Method: 1. B.S.D.W.

2. Treated with iron alum solution at room temperature overnight

3. Washed in running water for several minutes

4. Stained in the haematoxylin solution at room temperature overnight

5. Washed in water

6. Differentiate excess background dye in the iron alum solution differentiate in the borax-

ferricyanide solution if necessary.

7. Washed in water


Results: Myelin - Blue-black

MARALAND, GLEEN AND ZRIKSON'S A METHOD FOR AXONS IN

PARAFFIN SECTIONS (1954)

Preparation of staining solutions: a. 20% aqueous silver nitrate

b. Ammoniacal silver 20 ml ethanol was added to 30 ml 20% aqueous silver nitrate.

Conc. ammonia was added until the formed precipitate was almost dissolved. Two more

drops of ammonia were added

Method: 1. B.S.D.W.

2. It was placed in the 20% aqueous silver nitrate solution at 37°C for 30 min

3. Treated with 10% formalin for 15 min

4. Drained the slide and treated with ammoniacal silver solution for 30 seconds

5. Washed with 10% formalin for 1 min

6. Washed in water

7. Toned in 0.2% gold chloride for 2 min

8. Washed in water

9. Fixed in 5% hypo for 5 min

10. Washed in water

11. D.C.M.

Results: Nerve fibres, Neurofibrils, Neurons, R.B.C. Black Backgroun Grey

Notes: Sections are better celloidinized using 1% celloidin to give a thick protective coat. This not

just to keep the. section on the slide, but also when the celloidin coat is removed at the

conclusion of the technique much of the background silver precipitate will be removed

Toning helps suppressing a heavy background, but a clear distinction between collagen and

nervous elements cannot be made Glass slides should be carefully cleaned before staining to

reduce non-specific precipitation and silvering

Repeated step (4) & (5) is possible if under-impregnation

O.P.G METHOD FOR CALLS OF ANTERIOR PITUITARY (SILDDARS

1961)

Preparation of staining solutions: a. Orange G Orange G 500 mg Phosphotungstic acid 2 g Absolute alcohol 95 ml Distilled

water 5 ml

b. Acid Fuchsin Acid fuchsin 500 mg Glacial acetic acid 0.5 ml Distilled water 99.5 ml

Method: 1. B.S.D.W.

2. Stained nuclei in celestine blue-haemalum for 5 min

3. Differentiate and blue

4. Washed in water,then 95% alcohol

5. Stained with orange G for 10 min

6. Rinsed in distilled water

7. Stained in acid fuchsin solution for 2 min

8. rinsed in distilled water

9. Treated with 1% aqueous P.T.A.

10. Rinsed in distilled water

12. Stained in light green for 1 min

13. Washed in water

14. D.C.M.

Results: Nuclei Blue-black Basophil cells Red-purple Acidophil cells, R.B.C. Yellow Connective tissue

Green

Notes: - the staining of the basophils by acid fuchsin is progressive and should stain until the cells

are a well defined red-purple

- this is one of the trichrome staining and the mechanism is similar to trichrome technique

- normally, in the pituitary over 50% of the cells are termed as chromophobe cells, that is

the unstainable cells and are thought to be the cells for secretion of adrenocorticotrophic

hormone (ATCH). 40% of the rest are called acidophils ((X) , they are usually clamped

together in the lateral wings of the anterior lobe. The remaining 10% are termed as

basophils (B) , they are usu. larger than acidophils and are divided into two groups,

basophil-S-cells and basophil-R-cells

TRIPAS ( PAS-ORANGZ G METHOD )

Preparation of staining solutions: a. 1% periodic acid

b. Schiff's reagent

c. 2% orange G in 5% phosphotungstic acid

Method: 1. B.S.D.W.

2. Oxidized in periodic acid for 5 min

3. Washed in running water for 5 min and rinsed in distilled water

4. Placed in Schiff's reagent for 5 min

5. Washed in running tap water for 5-10 min

6. Counterstained nuclei lightly with Hx

7. Differentiate and blue

8. Washed in water

9. Stained in orange G for 20 seconds

10. Differentiate in tap water until macroscopically the section is pale yellow. Check

microscopically that only the R.B.C.'s and acidophi cells are yellow

11. D.C.M.

Results: Basophil cells - Magenta Acidophil cells - yellow R.B.C. - Yellow Nuclei - Blue-black

Chromophobes - Pale blue-black Nuclei - Red

Notes: Other structures, e.g. collagen, will be PAS positive. If this obscures the delicate staining, the

oxidation and Schiff,s staining times should be reduced Nuclei - Red

Methyl violet/crystal violet method (after Jurgens 1875)

Preparation of staining solutions: a. 1% aqueous methyl or crystal violet solution

b. 0.5% acetic acid

Method: 1. B.S.D.W.

2. Stain in 1% aqueous methyl or crystal violet for 2-5 min.

3. Wash in water and examine with the microscope.

4. Differentiate in 0.5-1% acetic acid until amyloid is purplish-red or red in good contrast with

the blue-violet staining of nuclei and normal tissue.

5. Wash in running tap water for at least 5 min.

6. Drain the slide and while the section is still moist, mount with modified Apathy's gum syrup.

Ring coverslip with nail varnish.

Results: Amyloid purplish-red to red Nuclei, cytoplasm, and connective tissue shades of blue-violet

Documents

Procedures of Histochemical