Preparation of tissue for microscopy

Microscopy dates from the 17th century when Hooke and Malpighi employed simple lenses in the study of various structural features. Between 1673 and 1716, Leeuwenhoek developed compound lenses, and by the early 19th century, the compound microscope had become highly sophisticated.

Usefulness of any type of microscope depends not only upon its ability to magnify but also upon its ability to resolve details. Beyond certain limits, magnification adds no new details. The useful magnification of an ordinary light microscope is about 1500x. Resolving power is a measure of the capacity of the microscope to clearly separate two points close together. Beyond the resolving power of any microscope two points will appear as one. The resolution with lens system is limited by the wave length of light and by numerical aperture or light gathering capacity of an objective lens. The resolving power of best constructed light microscope is about 0.2 micron.

Preparation of tissues:

Cells, tissues and organs can not be studied properly unless they are suitably prepared for microscopic examination. Two methods are used for tissues preparation

•     Methods involving direct observation of living cells.

•     Methods employed with dead cells.

Living cells are usually more difficult to handle & are available for a short period only. But it is important that one should be aware of the methods by which living cells may be observed and understand the ways in which they differ from fixed cells. In living cells, structure and functions may be studied simultaneously. Living cells can be seen moving, ingesting foreign materials, occasionally dividing, and carrying on other functions.

Observation of Living Tissues:

Unicellular organisms and occasionally free cells may be studied directly under microscope while they are still alive. Free cells are colorless and structures within them lack contrast. This difficulty may be overcome by using phase contrast microscope. Human blood cells are easily obtained and can be studied in films while surrounded by plasma, their natural environment.

Prolonged preservation of living cells outside the body can be achieved by a technique known as tissue culture. Fragments of tissues are removed aseptically, transferred to a physiological medium and kept at a temperature normal for the animal from which the tissues were taken. The culture is placed in thin glass vessel or on a cover glass mounted over a hollow glass slide. In this way they are available for observation under microscope. In such culture, growth, multiplication and in some cases, differentiation of cells into other cells type can be observed directly. Tissue culture is a valuable method for the study of cancer and the activity of many viruses.

Two staining methods have been applied successfully to living animals or to surviving cells.

Vital Staining:

In this case dyes are injected into living animals. The activity of certain cells will result in the selective absorption of the coloring materials by these cells e.g. staining of macrophages by trypan blue, on the basis of their ability to phagocytize foreign particles.

Surpravital Staining:

It involves the addition of a dye to a medium of cells already removed from the organism e.g. staining of mitochondria in living cells by Janus Green, of lysosome by neutral red and nerve fibers and cells by methylene blue.

Preparation of dead tissues:

The most convenient way to study histology is to use sections, each of which is more or less a permanent preparation. A section is prepared by cutting a thin slice from a small piece of fixed tissue, which is then stained, mounted in a medium of suitable refractive index on a slide and finally covered with a cover slip. The preparation of histological sections, involve the following procedure:

Removal of specimen:

For best histological preparation, the material should be removed from an anaesthetized animal or immediately after the death of an animal. In case of human material, this is scarcely possible. Surgical material represent the best source of human tissue since frequently, some normal tissue is removed together with an abnormal or diseased tissue.


Its primary objective is to preserve protoplasm with the least alteration from the living state. Fixative fluids acts to preserve protoplasm, inhibit autolytic changes and bacterial growth. Thus, it must be performed promptly to avoid tissue digestion by enzymes present in the tissues (autolysis).

Most fixing fluids coagulate protoplasm, thus rendering it insoluble and harden the tissue, so that sectioning is facilitated. They may or may not preserve carbohydrates and lipids. Many fixatives also increase the affinity of protoplasm for certain stains. Most commonly used fixative agents are formalin, alcohol, and certain acids i.e. picric acid, acetic acid and osmic acid. No single fixative possesses all the desirable qualities and many reagents are used in mixtures such as Bouin’s fluid containing picric acid, formalin and acetic acid and Zenker’s fluid composed of formalin, potassium dichromate and mercuric bichloride. The choice of fixative is usually determined by the particular tissue or compound that is to be studied and by the staining method to be used.


Its purpose is to provide rigid support to the tissue blocks so that they may be cut into thin sections. Prior to embedding, the fixed tissues is washed, to remove excess fixative and then dehydrated by passing it through increasing strength of alcohol or other dehydrating agents. This tissue is then cleared. This process involves the removal of dehydrating agent and its replacement by some fluid which is miscible both with the dehydrating agent and with the embedding material. Clearing agent include xylol, chloroform, benzene and cedarwood oil. After clearing, the tissue is infiltrated with embedding agent, usually melted paraffin or celloidin. After infiltration, the embedding agent is made to solidify so that a firm homogenous mass containing the embedded tissue is obtained.


Tissue embedded in paraffin may be sliced very thin. For the majority of microscopic work, sections are between 3 and 10 um thick. To cut such sections a microtome is used. Each section is transferred to a clean glass microscopic slide, on which a little egg albumin has been smeared. Water is run under the section and slide placed on a warming stage. The water evaporates and the section settles down onto the glass surface to which it becomes attached. The mounted section is now ready for staining.


The purpose of staining is to enhance natural contrast and to make cells and tissue components and extrinsic material more evident. Most stains are employed in aqueous solution and thus to stain a paraffin section, it is necessary to remove the paraffin solvent or decorating agent, usually by xylol. This step is omitted in the case of a section which has been embedded in celloidin. The section is then passed through descending strengths of alcohol prior to staining.


After staining excess dye is removed by washing with water or alcohol depending upon the solvent of the dye and the section is dehydrated through ascending grades of alcohol. Following absolute alcohol the section is transferred to a section of clearing agent. After removal from the cleaning agent, a drop of mounting medium e.g. Canada balsam which has a refractive index similar to that of glass, is placed on the section and allowed to dry.