How to Prepare a Live Specimen for Microscopic Study ? | Zoology

Get the answer of: How to prepare a live specimen for microscopic study.

A. Whole Mount Preparation:

Killing and fixing:

In case of live specimens they are to be killed. Killing must be instantaneous. This is done with chloroform, ether, acetone, alcohol, formalin, menthol, Bouin’s fluid, etc. These chemicals serve as fixatives also. Specimens contracting or retracting suddenly in contact with chemicals are narcotized with menthol, thymol, poisonous gas, etc., and then killed.

Staining:

Staining is the act of giving colour to some­thing. The killed or preserved specimens are thoroughly washed with water. This is followed by staining. In case of aqueous dyes, the speci­mens are first stained and then dehydrated. Dehydration means removal of water. In the use of alcoholic dyes the staining is done after dehydration to that concentration in which the dye is dissolved.

The period required to stain a specimen depends both on the size of the object and concentration of the dye in the solution. Over-staining, if any, is corrected by treatment with extremely diluted acid solution in water or alcohol according to the case, for a short period followed by immediate washing with water or alcohol. The water is removed from the speci­men in steps by passing it through 50%, 70%, 90% and 100% alcohol.

Permanent mounting:

The dehydrated specimen is cleared in clove oil. The oil is washed in xylol and the specimen is mounted on a slide in Canada balsam or DPX (Distrene Plasticiser xylene), and covered with a cover slip. Grooved slides are used for thick specimens.

1. Clean the slide and cover glass with a neat cloth.

2. Put a drop of mounting medium (Canada balsam or DPX) in the centre of the glass slide with a glass rod (the refractive index of Canada balsam and DPX being same as that of glass slide).

3. Transfer the tissue or organism from the clearing agent (xylol) on the slide by a brush or forceps.

4. Take a cover slip and place its one edge in the side touching the drop of mountant and hold its other edge obliquely either with a needle or forceps.

5. Gently lower down the needle till the cover slip touches the mountant and covers the materials (Fig. 22.1).

6. After mounting, the slide may be dried in hot plate or chamber, at 40-50°C.

7. The excess Canada balsam or DPX com­ing out around the cover slip should be re­moved with cotton wool soaked in xylol.

Labelling:

After permanent preparation the slide should be properly labelled. On one side of the slide the name of the material, date of collection, place of collection, etc., should be written with black Indian ink.

Temporary mounting:

Specimens do not need dehydration or clearing. Mount in water or saline solution or in 50% glycerine. If required the cover slip is sealed with bees wax or nail polish. A turn table is very convenient for the purpose.

B. Histological Preparation:

Collection of tissue:

For sectioning, tissues are to be collected from live specimens. In small vertebrates, the animal is paralyzed by damaging the brain. In larger vertebrates, the same result is achieved by striking the head against a hard object.

In still large species the tissues are collected immediately after killing the animal for some other purpose or a small piece of tissue is cut by surgical operation. In invertebrates, the tissues are usually collected from live speci­mens without such pre-treatment.

Objects to be studied under a microscope need preparation. If the object is small enough, through which light can pass, and the study in finest details is not required a whole mount preparation will be sufficient. In case of large objects they are to be cut in thin sections for the study. This is called histological preparation.

Fixing:

Killing the tissue without any structural or chemical change is known as fixing. In prac­tice, it has not yet been possible to prepare a fixative which satisfies all the requirements fully. The tissues are cut into small pieces, washed with physiological solution, if required, immersed in the fixative and left there for a definite period.

The common fixatives used are Bouin’s fluid (aqueous or alcoholic), Zenker’s fluid, Carnoy, Carnoy-labrum, etc. The specimen tubes with the fixative and tissues should be subjected to a vacuum pump for a few minutes to remove any air in the tissue which may enter during collection. Air inter­feres with the processing. The time required for fixation varies with the tissue.

Washing:

This is usually done with water till the reagent is completely removed. If the per cent of water in the fixative is below 50, it should be washed only in solution recommended for the purpose.

Dehydration:

Carry the tissues through grades of alcohol, viz., 50%, 70%, 90% and 100%, the period of treatment with each grade depending on the size of the tissue. Give two changes in absolute alcohol.

Clearing:

Transfer the tissue to ceder wood oil. Within 24 hours the tissues turn transparent and are ready for infiltration.

Infiltration:

This is to replace oil with paraffin. The intercellular spaces are completely occupied by paraffin. The tissue is washed in xylol and trans­ferred to a covered porcelain crucible contain­ing xylol and molten paraffin at a ratio 1: 1 and the crucible is put in a temperature controlled paraffin bath for 10 to 15 minutes. After the period, the tissue is transferred to another covered crucible containing molten paraffin only, kept in the same bath.

The time required for infiltration varies with tissues. Usually infil­tration is completed within 90 to 120 minutes. After the desired period paraffin blocks are made with the tissues. Paraffin’s of different melting points are used in different seasons.

Block-making procedure:

Pour molten paraffin in a watch glass or in a block making mould and put the tissue in the middle of it. After hardening, tream the extra paraffin to make a rectangular block keeping the tissue in the middle.

Section cutting:

This is done with a microtome. Sections for routine work are usually cut 5 to 6 μm thick. A grease-free slide is taken, one surface of the slide is marked with a diamond pencil. A drop­let of Mayer’s albumen is smeared on the marked surface.

A thin film of water is put on the slide and the paraffin ribbon with sections are placed on it. The slide is heated on a hot plate and the sections are properly stretched. The water is drained off and the slide is dried on a hot plate.

Staining of Histological Slides:

Sections of tissues are usually stained with two dyes to bring contrast between different histological structures. This makes detailed study easier. Staining with two dyes is known as double staining. The most common double staining practiced in class work is with haematoxylin and eosin.

The haematoxylin being basic dye imparts blue colour to acidic materials, viz., nucleic acids, which are con­centrated in the nucleus and only scattered in the cytoplasm. The eosin is an acid dye and the cytoplasmic materials being basic in nature are stained by it. The result is, the nucleus and only a small fraction of the cytoplasm appear blue, while the rest of the cell takes up red colour.

Technique:

The slide with the paraffin ribbon contain­ing sections of tissue is dipped in xylol for about 5 minutes. The paraffin dissolves. The slide is transferred to absolute alcohol. Haematoxylin is in aqueous medium.

Presuming eosin is in 90% alcohol the procedure to be followed is given below:

Time in each grade of alcohol is about two minutes with two changes in absolute alcohol. Time required for staining and differentiation is determined by trial. A small amount of Canada balsam or DPX is put on the slide, depending on the size of the cover slip to be used.

The amount should be just enough to form a very thin film on the slide. The cover slip is put on it. If air bubbles are locked between the cover slip and the slide, it may be removed by leaving the slide overnight on a hot plate.

Staining procedure:

Double Staining (Summary)