Technique Of Microtomy


Dissect open the animal and collect the tissues in mammalian Ringer Solution in a petri dish. Cut the tissues into small pieces. Flush the tissues like stomach, intestine etc. with saline water using a dropper. Press the glandular tissues like seminal vesicles, prostate gland etc. between two thick filter papers to remove the oily secretions.

Cut the tissues in pieces as you would like to see them in paraffin blocks according to the size and length. Fix the tissues separately in aqueous Bouin’s fluid (Saturated aqueous solution of picric acid—75 ml + formalin 40%–25 ml + glacial acetic acid—5 ml). Fixation time is 10-24 hours. Wash the tissues thoroughly in water to remove traces of picric acid so that it may not interfere in staining and in the process of dehydration.

DIFFERENT STAINS

1. Delafield’s haematoxylin:

Haematoxylin crystals—4 grams

Absolute alcohol—25 ml

Saturated solution of ammonia alum—400 ml

Glycerin—100 ml

Methyl alcohol—100 ml

Dissolve 4 grams of haematoxylin crystals in 25 ml of absolute alcohol and add to it 400 ml of saturated solution of ammonia alum and leave it exposed to sunlight and air in unstoppered bottle for about a week. Filter and add 100 ml of glycerin and 100 ml of methyl alcohol. Allow the solution to stand (uncorked) until the color is sufficiently dark and then filter.

2. Ehrlich’s haematoxylin

Haematoxylin crystals—2 grams

Absolute alcohol—100 ml

Glycerin—100 ml

Glacial acetic acid—10 ml

Distilled water—100 ml

Alum as above in excess

Dissolve the haematoxylin in absolute alcohol and add acetic acid and then glycerin and water. Let the mixture ripen in sunlight until it acquires a dark red color. Haematoxylin is used as a nuclear stain.

3. Eosin

Eosin powder—1 gram

Alcohol 90%–100 ml

4. Harris haematoxylin

Haematoxylin crystals—5grams

Absolute alcohol—50 ml

Ammonium aluminium sulphate or Potash alum—100 grams

Distilled water—1 litre

Mercuric oxide (Red)—2.5 grams

Dissolve haematoxylin in alcohol. Dissolve alum in water by heating. Remove from heat and mix the two solutions. Bring to boiling quickly. Remove from burner and slowly add mercuric oxide while cooling the flask under running water from the tap.

Advantage: This haematoxylin can be used after 24 hours, i.e. the next day and gives good results.

5. Mallory’s triple stain

Solution A. Acid fuchsin—1.0 gram

Distilled water—100 ml

Solution B. Aniline blue—0.5 gram

Orange G—2.0 grams

Phosphomolybdic acid—1.0 gram

Distilled water—100 ml

Solution C. Phosphomolybdic acid—1.0 gram

 Distilled water—100 ml

For Mallory’s triple stain these three solutions are prepared. In solution A sections are stained for 30 seconds, after which they are washed in distilled water for 2 minutes. Then stain in solution B for 1-5 minutes and wash in distilled water for 2 minutes. Then stain in solution C for 30 seconds and wash in distilled water for 1-2 minutes. Upgrade and mount in DPX.

Mallory’s triple stain is very good stain for animal tissues. You will enjoy the coloration of the cells.

STEPS FOR PROCESSING TISSUES

1. After fixing in Bouin’s fluid for 12-24 hours, wash well in tap water and then in distilled water. Remove as much of picric acid as possible by repeated washing.

2. Dehydrate the tissues in 30%, 50%, 70%, 90% and absolute alcohol, 4-6 hours in each grade. You can leave the tissue in 70% alcohol for any length of time and process a few pieces at a time.

3. Clear in xylene for 15-30 minutes. Small sized tissues are better as they become transparent quickly.  

Embedding in wax

Pour molten wax (50-60 0C range) in the bottle containing the tissues. Put in the incubator for 12-18 hours. Incubator should be set at 58-60 0C temperature, preferably at 60 0C. Keep filtered molten wax in beakers at the same melting range in the incubator. After infiltration of wax into the tissues for 12-18 hours make the tissue blocks.

Two L-shaped brass metallic pieces are put on the glass plate smeared with glycerin. Pour the molten wax into the cavity formed by connecting together the two metallic pieces and transfer the material in the middle of the wax-containing cavity. Remove the wax block after setting and drying. Trim the paraffin blocks to proper size and keep in paper covers to be used later.

Equipment required

1. Honning stone. Adair Dutt Co. Pvt. Ltd. India, 21, Asaf Ali Road, New Delhi-110001.

2. Leather-stropper with wooden base, from the same company.

3. Microtome machine. Westwox Optik model MT 1090. Precision Rotary Microtome.

4. Microtome knife. American Opticals from USA.

5. Machine oil. We use Usha tailoring machine oil, 3-in-one.

6. Haematoxylin stain for microscopy LOBA CHEMIE.

7. Eosin (soluble in alcohol) BDH or E. Merck Company.

8. Microslides, cover slips of different sizes.

9. DPX mountant.

10. Basic fuchsin (Para Rasanilene). Sigma chemical Co. USA.

Some hints while cutting sections

Cut at 7m thickness.

Prepare water bath at 58 or 60 0C.

Smear albumen (Mayer’s albumen:-White albumen, 50 ml; glycerin, 50 ml; sodium salicylate, 1.0 gram) on the slide. It will fix sections on the slide.

Put a few drops of 30% alcohol on the albumen smeared slide.

Cut the paraffin ribbon into bits containing 5 or 6 sections and put ribbon-bits on the slide with alcohol.

Put the ribbon into the water bath. The ribbon will completely stretch in the water bath without melting. Lift it with the same slide coated with Mayer’s albumen.

Rest the slide at an angle of 45 0C against a solid surface. Water under the material will get drained out in 6-8 minutes.

Put the slides in the incubator overnight (24 hours) at temperature of 56-58 0C.

Remove the slides next day and store them in a slide box as long as required.

Prepare the staining set in staining jars that contain grooves for holding slides. Six slides in 3 sets can be processed at a time. The procedure is as follows:

1. Downgrading or Hydration.  Xylene deparaffinization followed by 5 minutes each in absolute alcohol, 90%, 70%, 50% and 30% alcohol grades and distilled water.

2. Staining nuclei. In Harris haematoxylin 30-60 seconds depending on the tissues. Then dip the slides in acid water 0.5% (0.5 ml HCL in 100 ml of water) for one or two dips. Immediately put in tap water and wash for bluing of the nuclei in a glass trough with grooves.

3. Upgrading or dehydration. It is done by putting the slide for 5 minutes each in distilled water, 30%, 50%, 70% and 90% (2 changes) alcohol grades.

4. Cytoplasmic staining. Two dips in Eosin for 10 seconds each and wash for 3-5 minutes in 90% alcohol and then two changes of 5 minutes each in absolute alcohol. Then two changes of 5-10 minutes each in xylene for clearing. Then mount in DPX.

Keep the DPX mounted slides in the enamel tray overnight in the incubator for quick drying. Take out the slides next day and store in a slide box for examination under microscope at leisure.