2. Wash well in distilled water. 3. Place in a 5% aqueous solution of ammonium sulfide for 20–24 hours. 4. Wash in running tap water for 15–20 minutes, rinse well in distilled water and stain as desired.

 

Electrolytic Method

1. Decalcify in the electrolytic apparatus in formic acidhydrochloric acid for 1 to 4 hours.

 Electrolytic decalcifying solution

 Formic acid (90%) 100 cc

 Hydrochloric acid 80 cc

 Distilled water to make 1000 cc

2. Wash in running water for 24 hours.

3. Dehydrate, clear, and embed.

Processing of Tissues

Embedding in paraffin is accomplished most rapidly and

gives the best results when thin sections of soft tissues

are wanted. Since paraffin is not miscible with water, the

tissue must be dehydrated and then cleared in a solution

that is miscible with paraffin.

Automatic tissue processing units are usually employed

these days. Several manufacturers provide such units (Fig.

25.1). This instrument has several beakers, each filled

with a specific fluid in it. The basket containing the tissue

sections with identification tags is automatically dipped

and revolved in the beakers at preset timings. Usually,

paraffin-embedding process is started in the afternoon

and is complete by mornings on the following day. Various

schedules for paraffin processing are given below.

Method I

1. Alcohol 80% 1–2 hours

2. Alcohol 95%—2 changes 1–2 hours each

3. Alcohol absolute—3 changes 1–2 hours each

4. Xylene—2 changes 1–2 hours each

5. Melted paraffin—3 changes 1–2 hours each

6. Embed in paraffin and cool quickly.

Method II

1. Alcohol 80%—2 changes 1–2 hours each

2. Alcohol 95%—2 changes 1–2 hours each

3. Alcohol, absolute—3 changes 1–3 hours each

4. Chloroform—2 changes 1–2 hours each

5. Paraffin, melted—3 changes 1 hour each

6. Embed and cool quickly.

Method III

1. Alcohol 80%—2 changes 1–2 hours each

2. Alcohol 95%—2 changes 1–2 hours each

3. Alcohol, absolute—3 changes 1–2 hours each

4. Benzene—1 change 1–2 hours each

5. Paraffin bath—3 changes 1 hour each

6. Embed and cool immediately

 (In place of alcohol, one may use acetone. Xylene and

benzene can be used in place of one another. Benzene,

however, is carcinogenic and should be avoided).

FIG. 25.1: Automatic tissue processing unit

(Courtesy: Yorco Sales Pvt. Ltd)

Histopathology 793

Preparation of Sections

It is important that the knife used for cutting sections

be very sharp and without nicks. A perfect edge for a

microtome knife may be defined in simple terms as the

junction of two smooth plane surfaces at an angle of

about 14°. Knife sharpening may be done by mechanical

means on commercial automatic knife sharpeners or

done manually called honing and stropping. Nowadays

disposable knife blades with appropriate blade holders

are available.

Cutting Sections

After the paraffin block has been secured at the appropriate

place in the microtome, adjustment of the block and the

knife is now required. Keep a piece of cotton in a dish of tap

water and an ice cube in a petridish beside the microtome

(Fig. 25.2) at all times.

To facilitate sectioning, apply the wet cotton to the

surface of the block after rough cutting. Then place the ice

cube directly on the knife to flatten the side of the cube

that is to be applied to the surface of the block. Be sure to

crank the block back a fraction of a millimeter from the

knife edge, so that the first section cut after the block has

been soaked and chilled will not be too thick.

Collect the section ribbons in a bowl containing hot

water and unfurl or straighten the sections gently with a

fine tip camel’s hair brush.

Attaching Sections to Slides

The glass slides on which tissue sections are to be mounted

must be marked beforehand with the identifying case

number. A glass marking pencil is used for this purpose.

Paraffin sections may be attached to slides in several

ways. A small drop of Mayer’s egg albumin is smeared

over the surface of the slide with the finger and the excess

rubbed off with the heel of the hand, or it can be applied

with a clean foam rubber sponge. A sponge is usually

preferred so that the epithelial cells from the fingers will

not adhere to the slide and produce artefacts when stained.

Mayer’s Egg Albumin

Egg white 50 cc

Glycerin 50 cc

Mix well and filter through coarse filter paper, or through

several thicknesses of gauze. Add a crystal of thymol to

preserve.

Egg Adhesive from Dried Albumin

Albumin, dried 5.0 g

Sodium chloride 0.5 g

Distilled water 100.0 cc

Filter on Buchner’s funnel with vacuum. To 50 cc of filtrate,

and 50 cc glycerin, add a crystal of thymol to preserve.

Slides smeared by one of the above-mentioned egg

adhesives are taken below the floating section and the

section is placed in the center of the slide; leave it to dry to

be stained subsequently.

Technique for Frozen Sections

1. Fix small blocks of tissue in 10% formalin.

2. Wash blocks in water before freezing.

3. Put a drop of water on the holder and place the block

in position parallel to the knife edge.

4. Holding the block with the index finger or a glass slide,

turn on the coolant gas/fluid slightly. When block is

firmly fixed to holder, release more coolant gas/fluid

until the block is frozen.

5. Start sectioning and continue until a complete section

is obtained. Usually, the block will have thawed to

about right consistency by this time. If it is frozen

too hard, the sections may shatter. Allow the block to

thaw slightly and try again. If it has become too soft,

the sections will also shatter or fracture. The correct

temperature can only be judged by experience.

Sometimes rubbing the finger across the block will

give it the right consistency for good sections to cut. It

is best to cut slowly.

6. Lift the sections off the knife edge with the tip of the

little or ring finger which has been dipped in distilled

water. Place in a Petri dish of distilled water. Dry

the knife between sections as water will cause the

following section to be uneven or perforated.

7. Frozen sections may be stained with polychrome

methylene blue, hematoxylin and eosin, or fat stains.

FIG. 25.2: Rotary microtome

(Courtesy: Yorco Sales Pvt. Ltd)

794 Concise Book of Medical Laboratory Technology: Methods and Interpretations Staining

Staining may be done by the free flotation method or the

sections may be picked up on the slide first and stained as

usual.

Mounting

Fat stains must be mounted in glycerin jelly. Sections

stained by other techniques may also be mounted in

glycerin jelly or they may be dehydrated, cleared in xylene

and mounted in DPX mountant.

Removal of Pigments and Precipitates

Mercury Precipitate

If Zenker-fixed material stored for a long-time is to be

stained with alum hematoxylin, it will be found that areas

where the mercuric chloride is deposited stain deep

blue and distort the microscopic picture. Therefore, it is

necessary to:

1. Deparaffinize the sections by putting them on a hot

plate and then take through two changes of xylene,

absolute alcohol and 95% alcohol.

2. Place in alcoholic iodine (1 g iodine in 100 cc of 80%

alcohol) for 10 to 15 minutes.

3. Rinse in tap water.

4. Place in 5% aqueous sodium thiosulfate solution

(hypo) for 5 minutes.

5. Wash in running tap water for 10 to 20 minutes and

rinse well in distilled water before staining.

Formalin Produced Precipitate

Method I

1. Deparaffinize the sections through two changes each

of xylene, absolute alcohol, and 95% alcohol.

2. Rinse well in distilled water.

3. Let stand in saturated aqueous picric acid solution for

1 to 3 hours.

4. Wash well in running tap water.

Note

This picric acid solution will not bleach malarial pigment.

Method II

1. Deparaffinize the sections through two changes each

of xylene, absolute alcohol and 95% alcohol.

2. Rinse well in distilled water.

3. Place in bleaching solution for 5–10 minutes.

Bleaching solution

Hydrogen peroxide 25.0 cc

Acetone 25.0 cc

Ammonium hydroxide 1 drop.

4. Wash well in running tap water, and stain as desired.

Melanin Pigment

1. Deparaffinize the sections through two changes each

of xylene, absolute alcohol and 95% alcohol.

2. Rinse well in distilled water.

3. Place in 0.25% aqueous potassium permanganate

solution for 1 to 4 hours.

4. Wash well in water.

5. Place in a 5% aqueous oxalic acid solution or a

hydrobromic acid solution (HBr 1 part, distilled water

3 parts) until sections are clear (2–5 minutes).

6. Wash in running tap water for 10 minutes, rinse in

distilled water, and stain as desired.

Malarial Pigment

Method I

1. Deparaffinize the sections through two changes each

of xylene, absolute alcohol, and 95% alcohol.

2. Wash well in distilled water.

3. Place in a 5% aqueous solution of ammonium sulfide

for 20–24 hours.

4. Wash in running tap water for 15–20 minutes, rinse

well in distilled water and stain as desired.

Method II

1. Deparaffinize the sections through two changes each

of xylene, absolute alcohol, and 95% alcohol.

2. Place in a saturated alcoholic picric acid solution or 1

to 24 hours.

3. Wash well in running tap water and distilled water, and

stain as desired.