10% Formal Saline

10% FORMAL SALINE

Wear appropriate protective clothing eg. laboratory coat, gloves and goggles.
Weigh out 900g of sodium chloride and dissolve in warm water.
Unscrew cap on porthole of formalin mixing tank (beneath cut-up bench). Place a funnel in the front porthole of the formalin mixing tank and pour sodium chloride solution into the main tank. Replace screw cap on porthole.
Using the sealed pump attached to the formalin mixing tank , add 10 litres of formaldehyde to the main mixing tank.
Turn on the mains water tap (adjacent to the tank) and fill the formalin mixing tank to the 100 litre mark, using the line on the tank wall.
When mixed , using the sealed pump attached to the formalin mixing tank, pump 50 litres into the top tank where the solution is ready for use.
10% formal-saline solution can be dispensed from the tap with the red knob on the cut-up bench.

NB: See protocol on formaldehyde safety and COSHH form for formaldehyde for the hazards associated with this chemical.
100 litres of 10% Formal Calcium

900g sodium chloride
10L formaldehyde
90L water

4% PARAFORMALDEHYDE

4% PARAFORMALDEHYDE/PBS

1. Measure 100ml Phosphate Buffered Saline (PBS) into a measuring cylinder. Pour into the conical flask containing 4g of paraformaldehyde. Cover with parafilm and transfer to the fume hood: thoroughly shake - take care not to splash paraformaldehyde about - it is a rapid fixer and is TOXIC.

2. Place flask on top of the hotplate/stirrer inside the fume cupboard and set the heat control to 7 with moderate stirring. Allow the solution to warm up-it will turn from being cloudy to clear when ready. Inspect regularly to avoid over- heating and consequent spilling.

3. When the paraformaldehyde has dissolved, switch off the heat but leave to stir: do not handle for safety reasons. Allow to cool.

4. When cooled, transfer the fixative to a 4C refrigerator. Label appropriately and date.

0.4% PARAFORMALDEHYDE/PBS (FOR POST-DIGESTION FIXATION)

This fixative solution should be made up fresh.

Dissolve 0.4g paraformaldehyde/100ml DEPC-treated PBS in a conical flask and seal with Sealon film. On a heated stirrer (set to 6-7), inside the fume hood, mix until the milky solution turns colourless. Any cloudiness may be removed by adding a drop of 5M sodium hydroxide. Allow to cool before handling.

Safety

Paraformaldehyde is a rapid fixative and will cause harm if it comes into contact with the body.

WEAR GLOVES AND GOGGLES

ALCIAN BLUE TECHNIQUE

SECTIONS

4micron Paraffin wax sections

SOLUTIONS

Alcian Blue pH 3.1
Alcian Blue 1g
0.5% Acetic Acid 100ml
Alcian Blue pH 2.5
Alcian Blue 1g
3% Acetic Acid 100ml
Alcian Blue pH 1.0
Alcian Blue 1g
0.1M Hydrochloric Acid 100ml
Alcian Blue pH 0.2
Alcian Blue 1g
10% Sulphuric Acid 100ml
0.1% Nuclear fast red
Nuclear fast red 0.1g
Aluminium Sulphate 2.5g
Distilled water 100ml

METHOD

Take sections to water
Stain in alcian blue solution pH 2.5 for 5 minutes
Wash well in water
Counterstain in 0.1% nuclear fast red for 5 minutes
Wash in water
Dehydrate, clear . Mount sections in DPX

RESULTS

Mucins - Bright Blue
Nuclei - Red

NOTES

To further identify specific types of acid mucins you need to use the other pH solutions.
Strongly sulphated mucins - use pH 0.2 Alcian Blue
Weakly sulphated mucins - use pH 2.5 or pH 1.0 Alcian Blue
When using low pH solutions drain and blot dry instead of washing at stage (3) of the method.

ELASTIC VAN GIESON (EVG)

1. Bring sections to water.

2. Acidified potassium permanganate 2 minutes

3. Wash in water

4. 1% Oxalic acid 1 minute

5 Wash in water

6. Wash in 70% alcohol (optional)

7. Stain in Weigerts Resorcin Fuchsin at room temperature for 45 minutes

8. Wash in water 10 minutes

9. Differentiate in acid alcohol

10 Wash in water

11. Stain in Van Gieson for 5 minutes

12. Wash in water

13. Dehydrate, clear . Mount sections in DPX

N.B. Do not wash too long after staining in Van Gieson dehydrate and clear quickly

SOLUTIONS:

Weigert Resorcin Fuchsin

--See E.P.S.R. Method

Van Gieson (Unnas Variant)

--Acid Fuchsin 0.25g

--Nitric Acid 0.5ml

--Glycerin 10ml

--Picric Acid To saturation

--Distilled water 90ml

Acidified Potassium Permanganate

--0.5% Potassium permanganate 950ml

--3% Sulphuric acid 50ml

DECALCIFICATION

Many surgical specimens contain calcified areas which need to be decalcified before processing and sectioning. This is achieved as follows.

When the specimen is sufficiently fixed the selected tissue is placed into a labelled pot of 8% formic acid and left on the shelf in cut-up till the following day.
Each day thereafter the tissue is assessed until it is deemed soft enough to section after processing.
If it is not ready, the decalcifying fluid is changed and the pot is replaced onto the shelf for a further 24 hours.
This procedure is repeated daily until decalcification is complete.

Note: it is possible to test biochemically for calcium in solution to assess endpoint of decalcification. This is not done routinely.

FORMALDEHYDE PRECAUTIONS

Formaldehyde is toxic and the exposure limit is 2ppm for 10 minutes in every 8 hours. If you are affected by the vapour in any way e.g. eyes watering, leave the area immediately.
Containers containing formaldehyde solutions should only be opened in a designated ventillated area. Goggles , gloves, and an apron should be worn over laboratory coat.
Gloves, goggles and apron must be worn over laboratory coat when making up formol saline or changing the formalin on specimens. This procedure should be performed in a designated ventillated area.
Any spillages must be cleaned up immediately (see COSHH protocols)

HAEMATOXYLIN AND EOSIN TECHNIQUE

SOLUTIONS

(i) Gills Haematoxylin

Haematoxylin 6.0g
Alluminium Sulphate 4.2g
Citric Acid 1.4g
Sodium Iodate 0.6g
Ethylene Glycol 269ml
Distilled Water 680ml

(ii) Eosin

Eosin Yellowish 1.0g
Distilled Water 100ml

(iii) Lithium Carbonate 1%

Lithium Carbonate 1g
Distilled Water 100g

(iv) Acid Alcohol 1%

70% Alcohol 99ml
conc. Hydrochloric Acid 1ml

(v)Scott's tap water

In a beaker containing 1L distilled water, add 20g sodium bicarbonate and 3.5g magnesium sulphate. add a magnetic stirrer and mix thoroughly to dissolve the salts. Using a filter funnel, pour the solution into a labelled bottle

METHOD

Take sections to water.
Place sections in haematoxylin for 5 minutes.
Wash in tap water.
'Blue' sections in lithium carbonate or Scott's tap water.
Wash in tap water.
Place sections in 1% acid alcohol for a few seconds.
Wash in tap water.
Place sections in eosin for 5 minutes.
Wash in tap water.
Dehydrate, clear . Mount sections in DPX

RESULTS

Nuclei Blue-black
Cytoplasm Varying shades of pink
Muscle fibres Deep pinky red.
Fibrin Deep pink
Red blood cells Orange/red

GRAM STAIN

SECTIONS

4µm paraffin wax sections

SOLUTIONS

1. Hucker-conn ammonium oxalate - Crystal violet

Crystal violet 2g
95% Alcohol 20cm3
Ammonium oxalate 0.8g
Distilled water 80cm3

Dissolve the crystal violet in the alcohol, and the ammonium oxalate in the distilled water, and mix the 2 solutions. Stable for about 2 years but may need occasional filtering.

2. Weigerts Iodine

Potassium iodide 2g
Iodine crystals 1g
Distilled water 100ml

Dissolve the potassium iodide in 2-3mls of the distilled water and then dissolve the iodine crystals. Dilute to 100mls in the rest of the distilled water.

3. 0.1% Nuclear fast red

Nuclear fast red 0.1g
Aluminum sulphate 2.5g
Distilled water 100mls

METHOD

Take sections to water
Stain in filtered crystal violet for 2 minutes
Rinse in water
Treat with Weigerts iodine for 2 minutes
Rinse in water briefly
Differentiate in acetone until the section is colourless
Rinse briefly in water
Counterstain in nuclear fast red for 5 minutes
Dehydrate, clear . Mount sections in DPX

RESULTS

Gram positive organisms and some fibrin - Blue/black
Gram negative organisms and cell nuclei - Red

MASSON FONTANA

1. Sections to distilled water.

2. Treat with silver solution in a coplin jar in a water bath at 45ª for 30 minutes

3 Wash in water.

4. Counterstain in 1% eosin for 30 seconds.

5. Dehydrate, clear . Mount sections in DPX

RESULTS

--Melanin black

--Argentaffin,chromaffin and some lipofuscin black

--Background red

SOLUTIONS

SILVER SOLUTION

To 20 mls of 10% silver nitrate add concentrated ammonia drop by drop until the formed precipitate almost dissolves to the silver solution add 20 mls of distilled water then filter

SAFETY

Waste ammoniacal silver solution is potentially explosive. Collect all ammoniacal silver and put into the silver waste bottle - it contains saturated sodium chloride to precipitate out the silver, this not only makes it safe but allows the silver to be sold for reclamation. Do not pour waste solutions down the drain.

MASSONS TRICHOME

SECTIONS

4µm paraffin wax sections

SOLUTIONS

1. WEIGERTS IRON HAEMATOXYLIN

Haematoxylin solution
Haematoxylin 1g
Absolute alcohol 100mls

Allow to ripen naturally for 4 weeks before use

Iron solution
30% aqueous ferric chloride
Concentrated hydrochloric acid 1ml
Distilled water 95mls

This solution is filtered and added to an equal volume of the haematoxylin solution immediately before use. The mixture should be a violet/black colour and must be discarded if it is brown.

2. PONCEAU - ACID FUCHSIN SOLUTION

Equal volume s of 0.5% ponceau 2 R in 1% acetic acid and 0.5% acid fuchsin in 1% acetic acid.

3. 1% phosphomolybdic acid

4. 2% light green in 2% citric acid diluted 1:10 with distilled water prior to use

5. Aniline Blue

Boil 97.5mls distilled water and add 2g Aniline Blue while still hot. Then add 2.5mls glacial acetic acid, cool and filter

METHOD

Take sections to water
Stain nuclei with Weigerts iron haematoxylin for 10 minutes
Wash in water and 'blue' in lithium carbonate
Stain in 1% panceau-acetic acid solution for 5 minutes
Rinse rapidly in water
Differentiate in 1% phosphomolybdic acid for approximately 5 minutes
Drain and counterstain with light green or aniline blue
Dehydrate, clear . Mount sections in DPX

RESULTS

Nuclei - Blue/black
Muscle, red blood cells, fibrin - Red
Connective tissue - Blue or green

METHENAMINE SILVER FOR FUNGI

1. Bring sections to water.

2. Oxidise in 5% chromic acid for 1 hour.

3. Wash in running tap water.

4. Rinse well in distilled water.

5. Place in incubating solution at 60°c in the dark for approx 1 hour - needs checking microscopically. (Use coplin jar in a water bath and preheat solution before use).

6. Wash well in distilled water.

7. Tone in 0.1% gold chloride 2 mins.

8. Place sections in 3% sodium thiosulphate 2 mins.

9. Wash well in tap water.

10. Counterstain using sat alcoholic picric acid 1/2 hour.

11. Rinse in absolute alcohol.

12. Clear in xylene . Mount sections in DPX

RESULTS

--Fungal hyphae and yeast bodies black

--Background yellow

SOLUTIONS

5% Borax in distilled water.

Silver Solution

--5% Silver nitrate in distilled water 5ml

--3% methenamine (hexamine) in distilled water 100ml

Add the methenamine solution to the silver, a white precipitate forms, but clears on shaking. Both Solutions keep well at 4°C

Incubating solution ( make up fresh)

--Borax solution 5ml --Distilled water 25ml --Silver solution 25ml

OIL RED O

1. Cut 2 frozen sections.

2. Fix in 10% formal calcium.

3. Wash in water.

4. Rinse in 60% T.E.P. (triethyl phosphate).

5. Stain in filtered oil red o for 10-15 minutes.

6. Rinse in 60% T.E.P.

7. Rinse in water.

8. Stain nuclei with Mayers haematoxylin for 1 minute. (N.B. do not differentiate)

9. Blue in tap water.

10. Mount in aqueous mountant. (glycerin jelly).

RESULTS

--Lipid red --Nuclei blue/black

CONTROLS

--Not essential

--Fatty liver

--Negative control

NEGATIVE CONTROLS

Treat one section with a 50:50 acetone/xylene mixture for 5 minutes, wash in water then continue as from step 4.

PAPANICOLAOU METHOD

1. Fix slides in acetic/alcohol fixative for 15 minutes.

2. Absolute alcohol 2 minutes.

3. 70% alcohol 2 minutes.

4. 50% alcohol 2 minutes.

5. Tap water 2 minutes.

6. Stain in haematoxylin 4 minutes.

7. Rinse in tap water briefly.

8. Differentiate in acid alcohol 5 seconds.

9. Blue in tap water.

10. Dehydrate in absolute alcohol x2.

11. Stain in orange G 10 seconds.

12. Rinse in absolute alcohol x2.

13. Stain in E.A. 50 2 minutes.

14. Rinse in absolute alcohol x2.

15. Clear in xylene x3.

16. Mount sections in DPX

Acetic Alcohol Fixative.

Add 60 mls of glacial acetic acid to 100 mls of distilled water then add this solution to 2 l of absolute alcohol.

PERIODIC ACID SCHIFFS ORANGE G

1. Sections to water.

2. Treat with 1% periodic acid for 10 minutes.

3. Wash in water.

4. Stain in Schiffs reagent for 10 minutes.

5. Wash in tap water until sections appear pink.

6. Treat with 4% iron alum for 5 minutes.

7. Wash in distilled water.

8. Stain nuclei with Mayers haematoxylin for 5.minutes.

9. Differentiate in 1% acid alcohol.

10. Blue in tap water.

11. Stain with 0.2% orange g in alcoholic picric acid.

12. Dehydrate in absolute alcohol and examine microscopically.

13. Differentiate in 70% alcohol until the background appears colourless.

14. Dehydrate, clear . Mount sections in DPX

RESULTS

--Basophil cells magenta

--Acidophil cells yellow

--Red blood cells yellow

--Nuclei blue/black

--Chromophobes pale blue/grey

METHYL GREEN PYRONIN

1. Bring sections to water.

2. Rinse with Acetate Buffer pH 4.8.

3. Stain with the MGP solution 30 mins.

4. Wash with buffer and blot dry.

5. Rinse in equal parts acetone/xylene, then xylene.

6. Mount sections in DPX

RESULTS

--DNA (Nuclei) Green or Green/Blue

--RNA (Plasma cell cytoplasm, nissl substance, bacteria) Red

--Some mucins Red

SOLUTIONS

--2% Aqueous methyl green (extract with chloroform until methyl violet contaminant is removed) 9 ml

--2% Aqueous pyronin Y or G 4ml

--Glycerol 14ml

--pH 4.8 buffer 23ml

--MIX BEFORE USE

PERLS TECHNIQUE

SECTIONS

4µm paraffin wax sections

SOLUTIONS

1. INCUBATING SOLUTION

--2% Potassium ferrocyanide 10mls

--2% Hydrochloric acid 10mls

--Prepare fresh before use

2. 0.1% Nuclear fast red

METHOD

1. Take sections to water

2. Place in freshly prepared incubating solution for 10 minutes

3. Wash well in water

4. Counterstain in 0.1% nuclear fast red for 5 minutes

5. Wash in water

6. Dehydrate, clear . Mount sections in DPX

RESULTS

--Ferric iron - Blue

--Nuclei - Red

PERIODIC ACID, SCHIFFS

1. Bring sections to water.

2. Treat with 1% periodic acid for 10 mins.

3. Wash in water.

4. Treat with Schiffs reagent for 10 mins.

5. Wash well in tap water.

6. Stain nuclei with Carazzi haematoxylin for 2 mins.

7. Differentiate in acid alcohol.

8. Blue in Scotts tap water.

9. Dehydrate, clear . Mount sections in DPX

N.B. Do not over stain in schiffs reagent - it is irreversible. Wash for several mins at step 5 to bring out the colour of the Schiffs.

RESULTS

--Glycogen, other periodate reactive carbohydrates - magenta

--Nuclei- blue

SOLUTIONS

Schiffs reagent

1. Boil 200ml of distilled water, remove flask from bunsen and add 1g of basic fuchsin.

2. Allow to cool to 50°c.

3. Add 2g of potassium metabisulphate whilst mixing.

4. Cool to room temperature and add 2ml of concentrated hydrochloric acid mix and stand in the dark overnight.

5. Add a large amount of activated charcoal, shake well and filter. The solution should be clear/pale yellow.

Store at 4°c

REMOVAL OF FORMALIN PIGMENT

1. Sections to water.

2. Immerse in saturated alcoholic picric acid in a coplin jar - 20 mins.

3. Wash in water until all yellow colouration is removed from the section.

4. Proceed with staining method.

Immunofluorescence Staining

Direct Immunofluorescence Staining

Harvest and wash cells in Fluorescence Buffer (FB).
Add 50l of conjugated primary antibody (Ab) to cell pellet, vortex.
Incubate 20-30 min on ice
Wash 2x's with FB.
Resuspend cells in FB, or 1% paraformaldehyde, or 2g/ml propidium iodide buffer (PI).
Acquire and analyze on cytometer.

Indirect Immunofluorescence Staining

Harvest and wash cells in Fluorescence Buffer (FB).
Add 50l of unconjugated primary Ab to cell pellet, vortex.
Incubate 20-30 min on ice.
Wash 1x's with FB.
Add 50ml of secondary Ab , vortex.
Incubate 20-30 min on ice.
Wash 2x's with FACS buffer.
Resuspend cells in FB, 1% paraformaldehyde, or 2 g/ml propidium iodide buffer (PI).
Acquire and analyze on cytometer.

Reagents

Fluorescence Buffer (FB, 1 liter)

10x PBS / 100 ml
2.5% NBCS / 25 ml (new born calf serum)
0.02% NaN3 / 10 ml (2% NaN3 stock)
H2O / 865 ml

Filter before use. Store at 4° C.

Notes

Titer all Ab's and secondary reagents prior to use to determine optimal concentration.
Use appropriate isotype matched controls.
Airfuge Ab when necessary, i.e. if frozen.
Resuspend cells in a minimum of 250l.
Cell number may vary depnding upon experimental conditions, however 5 x 105 -1 x 106 cells is optimal.

Microsphere Quantitation Assay

Staining

Harvest assay.
After washing cells stain with antibody (Ab), according to standard immunofluorescence staining protocol.
Following final wash after staining, add constant volume (250-500 l) of 10% bead suspension in the absence or presence of propidum iodine (PI). Pipet accurately!

Acquisition of sample on cytometer

Acquire contents of tube on cytometer. It is important to threshold on SSC (12-16) to include the bead events.
Make sure beads and cells show up on light scatter and draw a gate around both populations. Exclude spurious events from debis or noise.
Acquire constant event# (beads + cells= 5000 to10000 events). Be sure to mix each tube before acquiring to assure good mixture of cells and beads.

Analysis of data

Analyze data to determine cell event # and bead event #. Set the following gates:
Gate 1: FSC vs SSC gate around beads
Gate 2: FSC vs SSC gate around cells
Calculate cell numbers:
# cells acquired X# beads added = #cells
# beads acquired tube tube
Example calculation:
2378 cells acquired X38400 beads added =45363 cells
2013 beads acquired tube tube
To find # beads added:
# of beads Xml beads =beads added
ml tube tube
Example calculation:
128 x 104 beads X0.030ml beads =beads added
ml tube tube

Reagents
Stock bead suspension

 Add 3-5 drops of 6.0 micron Polybead polystyrene Microspheres (Polysciences, Inc., Warrington ,PA) to 30-50ml FB. Store at 4°C.

 Calculate concentration of beads.

Notes

 Avoid aggregates in stock bead suspension, mix well before use.

 Gate 2 in the analysis section can be drawn to define any population cells based on multiple parameters.

References
1. Larson et al, J. Immunol. 151:138, 1993.
2. Dittel et al, J. Immunol. 154:58, 1995.
3. Pribyl et al, PNAS 93:10348, 1996.

Breast Cancer Reporting Protocol

Macroscopic Examination