Theory of Staining in histopathology

Theory of Staining in histopathology

Theory of Staining in histopathology

Tissues and their constituent cells are usually transparent and colorless when examined under the light microscope, with little or no differentiation of the various structures. Colouring, in other words, dyeing or staining of the sections of tissues makes it possible to see and study the physical features and relationships of the tissues and their constituent cells. It so happens that different tissues and indeed, different components of the cell, show differing affinities for most dyes or stains.


It follows therefore that no single staining method will demonstrate all the tissue structures present. It is often necessary to carry out several different staining methods on consecutive sections from a block of tissue in order to make a diagnosis Histochemistry is a special branch of histopathology that attempts to identify tissue and cell components with the results of specific chemical reactions which helps the trained pathologist or histotechnologist to establish the presence or absence of disease. Such specific strains have little or no affinity for another tissue elements. These are also often referred to as special stains. Examples are the demonstration of haemosiderin with Perlis Prussian blue reaction and of polysaccharides with periodic acid Schiff method.

A good number of biological scientists have put in a great deal of energy into the research of the great phenomenon of why certain stains have an affinity for some tissue structures and not for others. At this point in time, there is no single theory that has been universally accepted. It was the norm in

the past to support either the physical or chemical theory. But more and more workers now tend to base their acceptance on a combination of both the physical and chemical factors.

Physical Factors

The physical factors which influence staining reactions include: 

(1) Osmosis and capillarity These simple physical forces are deemed to be partly responsible for the penetration of stains into porous tissues. 

(ii) Absorption This factor is demonstrated by the action of certain stains on certain tissues in the presence of mineral salts. Sun light rays are selectively absorbed and cause colour to appear. 

(iii) Selective adsorption This factor is a principle which has been employed since long time by physical chemists. It is the characteristic of certain substances to adsorb certain ions from a solution more readily than from others. This factor is considered the most acceptable rationale of many staining reactions.

Chemical Factors The chemical theory is based on the assumption that certain parts of biological tissues are acid in nature, e.g., the nuclei of cells, while other parts, such as the cytoplasm, are basic in reaction. The colouring matter in basic dyes is contained in the basic part of the compound, while the acid radical

is colourless; and vice-versa for acid dyes. It therefore follows that acid tissue elements will have affinity for a basic stain while the basic structures have affinity for acid stains.

IMPREGNATION

Although the term staining is loosely used to describe the process of coloring tissue structure, there are some cell and tissue components that can be demonstrated not by staining but by a process of coating known as impregnation. Impregnation, unlike staining, makes use of the salts of heavy metals which are precipitated selectively on the cellular and tissue components. This technique has its greatest application in tissues from the central nervous system and for the demonstration of the reticulum. Impregnation differs from staining in that it comprises particulate precipitate. In the final analysis however, it displays many of the effects of true staining. Silver nitrate is the most commonly used compound for impregnation and can behave like a stain and present tissue elements in a nonparticulate union.

CHEMISTRY OF COLOUR IN DYESTUFFS

The chemical basis of color in dyestuff is made up of certain atomic units known as chromophores (color-bearers in Greek). When a chromophore is added to an uncolored molecule, it becomes a chromogen, which though colored, is not a dye. And to become a dye, the chromogen must consist of an organic compound with a chromophore in its structure and an ionizing radical, autochrome, as an additional side chain. The difference between a chromogen and a dye is that the colour of a chromogen is easily removed the colors are not "fast").

The autochrome acts as an electron donor to the chromogen. The main auxochromes are the -OH and-NH, groups. They help to impart the dye colour to the tissue because they are able to dissociate and form compounds. Sulphuric groups (-SO,H) and carboxyl groups (-COOH) may also function as auxochromes. The three main groups of chromophores are the

(1) Quinonoid ring, para or ortho (Fig. 6.1). The dyes include basic and acid fuchsin (Fig. 6.2). eosin, cystal violet, methylene blue, neutral red and natural dyes, e.g., haematein (Fig. 6.3).

Hydrogenation of chromophore results in the reduction of a coloured dye to form a colourless or *leuco' form. Schiff's reagent, though colourless, is not a typical leucobase because it is re-colourised, not by oxidation but on contact with an aldehyde.

CLASSIFICATION OF DYES

Dyes are classified in several ways. They can be grouped as natural or synthetic; or according to their reactions, tissue affinities and major applications.

NATURAL DYES

Haematoxylin Haematoxylin is a natural dye that is extracted from the core or heartwood of the tree Haematoxylin campechianum which is native to the region of Campeche in Mexico where its dye property had long been known to the natives. It was solely used in the textile industry to dye silks and wool until in 1862 when Waldeye firmly introduced it for use in histology. Today the tree is grown on a commercial scale mainly in Jamaica. The trees are felled at about 10 years, the bark and sapwood (outer layers) scrapped off, and the heartwood cut to about three feet logs for export. The logs are cut into chips for extraction.

The pure extract from the logs which is known as haematoxylin, has to be oxidised or ripened to its active form-haematein-for it to become an active dyestuff. During oxidation, haematoxylon loses two hydrogen atoms and takes on a quinonoid arrangement in one of its rings.

Oxidants Oxidation can be achieved either by exposure to air or sunlight, or by the use of chemical oxidants. Exposure to air is slow and may take up to 3-4 months, whereas the ripening is almost instantaneous when chemicals are used. The popular chemical oxidants are mercuric oxide, sodium iodate, potassium permanganate, hydrogen peroxide and calcium hypochlorite. Substances other than haematein are produced during oxidation though none of them is as effective as haematein. It is very important that the correct amount of oxidant is used in making up haematoxylin stains.

Mordants Haematein on its own has a poor affinity for tissues. It requires a mordant to render it effective. Mordants are substances that link the tissue and the stain and thereby bring about a staining reaction between them. Mordants used for haematoxylin staining are always divalent or trivalent salts or hydroxides of metals. They are believed to combine as hydroxides with the dye by displacing a hydrogen atom from it, and the remaining valencies serve to attach or link the stain--mordant complex to tissue components, and bring about a staining reaction (Fig. 6.7).

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The stain-mordant complex is known as 'lake'. The commonly used mordants are salts of aluminium, iron, chromium and copper. The alum salts remain the most popular simply because they were most readily available in the early days and so traditionally, histologists still stick to them.

Mordants need not necessarily be incorporated in the stain. Some fixatives such as mercuric chloride or picric acid act as mordants for Mallory's phospho molybdic and phosphotungstic acid methods for connective tissue. This is called premor-danting. Also, when myelinated fibres of the CNS are to be demonstrated, the tissue is usually fixed in 10 % formal saline and prior toembedding, it is mordanted in Weigert's primary mordant which is a mixture of potassium dichromate and a flouroch-rome.

Cochineal

One of the oldest dyes used in histology is Cochineal. It is a dye extracted from the bodies of female cochineal insects. When obtained, the dye is treated with alum and the result is a product relatively free from extraneous matter and is known as carmine. Carmine is widely used for staining zoological specimens. When combined with picric acid it is known as picrocarmine and is very useful in neuropathological work. It stains the nuclei very well and is used for the demonstration of glycogen as in Best's carmine, and mucin as in Southgate's mucin carmine staining method.

Orcein

One of the few vegetable dyes in histology is the Orcein. It is extracted from lichens by the action of ammonia and air. It is violet in colour, soluble in alkalis and weak acids. It is employed in the Taenzer-Unna orcein method for the demonstration of elastic fibres. A synthetic brand is commercially available.

Litmus

Like the orcein, litmus is extracted from lichens and is exposed to ammonia and air but in addition,

it is treated with lime and potash or soda. Litmus was greatly used as an indicator but is now superseded by synthetic brands.

Saffron

Saffron as a dye has only one use in histology. It has been incorporated by Masson into connective tissue stain. It is extracted from the plant, Crocus sativus.

SYNTHETIC DYES

Nearly all the staining agents employed in histology are synthetic aromatic dyes made from coal tar derivatives. In fact they were once referred to as coal tar dyes. A few exceptions are the inorganic substances that are used for staining or impregnation, e.g., silver nitrate, gold chloride, iodine, osmium tetroxide and potassium permanganate.

Synthetic dyes are derivatives of the hydrocarbop benzene (CH) which is represented by a benzene ring as in Fig. 6.8. When the C and H atoms are omitted, it can be drawn as Fig. 6.9. Compounds such as benzene have absorption band in the ultraviolet range of the spectrum.

When combined with a chromophore, the benzene becomes coloured and visible (chromogen), as seen in Fig. 6.10. When an auxochrome, e.g., a hydroxyl group displaces another hydrogen atom, a dye compound (picric acid) is formed (Fig. 6.6). A synthetic dye therefore is a hydrocarbon (benzene) derivative to which a chromophore and an auxochrome groups have been added.

BASIC, ACID AND NEUTRAL DYES

Ordinarily, the nature of the autochrome usually determines whether a particular dye is acid or basic in reaction. 

Basic dyes/stains In basic dyes, the colouring substance of the compound is a base and the colourless acid radicle, usually derived from hydrochloric, sulphuric and acetic acids, is combined with the basic substance; e.g., basic fuchsin is the chloride salt of the base rosaniline (Fig. 6.2a). The acidic cell structures, e.g., nuclei, have an affinity for the basic ions of the stain and so are regarded as basophilic. 

Acid dyes/ stains in these stains, the acidic radical is the active or colouring agent while the basic part is inactive. For example, acid fuchsin is composed of the sodium salt of a sulphonate of rosaniline (Fig. 6.2b). Basic cell structures, e.g., cytoplasm, have an affinity for the acidic ions and so are regarded as acidophilic. Eosin is an example of an acid dye/stain.

Neutral dyes These are usually mixtures of basic and acid dyes, and which are not easily soluble in water but readily so in alcohols. They are made up of large molecular complexes. Basic components of the dye stain the nuclei while the acid parts stain the cytoplasm. Thus neutral dyes stain both the nuclei and cytoplasm. The Romanowsky stains (e.g., Leishman stain) are examples of neutral dyes.

Selective solubility The mechanism of selective solubility is based on the fact that certain colouring agents are more soluble in some tissue elements than in others. For example, oil red 0 selectively stains and dissolves in neutral fat but does not enter other tissue components.

STORAGE AND MAINTENANCE OF DYES

Dyes are generally supplied in solid-state. They are better stored in a cool dark place. Usually, the quality of the dye is taken for granted based on the integrity of the supplier or manufacturer. But for the avoidance of doubt, some laboratories carry out quality control tests on new and very old stocks.

In making up the stains, distilled water is preferred.
Most importantly the manufacturer's directions should be adhered to as much as possible. Pure chemicals should be used in the formulation of the dyes. Stains are best stored in glass bottles or flasks and are kept in a cool place, properly labelled with the name and the date of preparation. In some cases filtering is required before use.

Dyes and stains are known by several names. It is therefore essential to quote the colour index number or lot number of any particular dye when ordering. Table 6.1 shows the colour index (CI) number and solubility table of some common histological stains.

STAINING PROPERTIES OF DYES

The staining property of a dye may be microanatomical or cytological. Microanatomical stains are those used to demonstrate the general relationship of tissues to each other. The stains clearly differentiate the cytoplasm from the nuclei while the other cellular structures are not well highlighted. Cytological stains, on the other hand, are used for demonstrating the tiny structures in the nucleus and the cytoplasm and not the tissue types.

Indirect staining When a mordant is employed to help bring about a staining reaction, the method is referred to as indirect staining method, e.g., haematoxylin and eosin staining method. 

Direct staining When a mordant is not necessary for staining reaction to occur, this is direct staining. Most aqueous or alcoholic aniline stains are in this group. 

Accentuators These are chemical substances that increase the colouring power, crispness and selectivity of a stain. Unlike mordants, they do not bind or link the stain to the tissue. It is claimed that some of these chemicals act by reducing the surface tension and others act as chemicophysical catalysts.
Accentuators are also known as accelerators especially when they are used in conjunction with certain hypnotic drugs such as barbital (Veronal), and chloral hydrate during the metallic impregnation of nerve fibres. Examples of accentuators are phenol in carbol fuchsin, aniline used in gentian violet and potassium hydroxide in Loefflers's methylene blue.

Progressive staining Stains which colour the tissue components in a specific order are called progressive stains. 

Regressive staining This is when tissues are overstained and the excess stain is removed selectively (differentiation) to give the desired intensity.

Vital staining When inclusions of live cells or tissues are stained, it is called vital staining. 

Supravital staining When the living cells are stained after being removed from the body, it is called supravital staining. 

Intravital staining This refers to the staining of the cells while still part of the body. 

Metachromasia Most dyestuffs stain tissues or organisms in various shades of their own colour. Some dyes somehow stain certain tissues in a colour different from their own colour and that produced in other parts of the tissue. This phenomenon is known as metachromasia and is seen only in basic dyes of the aniline type. Most of the dyes belong to the thiazine group, others may be of triphenylmethanes or azo types. Examples of metachromatic stains are New methylene blue,

toluidine blue, crystal violet, methyl violet and thionin. But the commonly used ones are the toluidine blue, methylene blue and thionin.

This phenomenon depends on (1) the dye and (2) the nature of the tissue parts that will pick up the dye and show metachromasia. The original colour of the dye is as a result of the single molecular form in solution. These dyes do polymerise, and as more and more polymers (dimers and trimers) are formed, the solution changes colour. For example, toluidine blue in the simple molecular form in solution is blue. When the dimers and trimers increase in number, it turns violet and eventually the full red metachromic colour appears. This is due to the polymers of the basic dye molecule. The dyes are cationic, and it is believed that all tissue components that exhibit metachromasia are composed of large anionic molecules that contain many sulphate, phosphate or carboxylic acid radicals.

Note

1. Use 0.1% or less concentration of the dye in 30% ethanol or as 0.1% to 1% aqueous solution.

2. Do not overstain in order not to lose the metachromatic effects, for most methods staining time is 1/2-1 minute.

3. Beware of impurities that are present in some dyes (, e.g., thionin) as these may stain different colours which can be mistaken for metachromasia.

Negative staining This is the method employed to examine bacterial morphology and demonstrate any envelope or capsule around the organism. The organism and the staining agent are mixed on a slide, and examined microscopically. The unstained structures are sharply seen against a dark background.

STAINING EQUIPMENTS AND MATERIALS

Staining of histology slides is generally done on a specially designated bench with an appropriate sink. The sink should be shallow, about 15 cm deep; and should have an efficient overflow. A slide washing tray, usually of stainless steel, may be placed in the sink. The sink must be kept clean.

Within the staining area, there should be a bunsen burner to heat up stains, a thermostatically controlled hot plate for melting wax around the sections and for hardening mounting media, a 37°C incubator and a 60°C oven. There should also be a microscope for controlling the degree of staining reaction.

The histology slides may be stained in any of the following ways:

(i)using staining dishes

(ii) using staining racks

(iii) using a staining machine

Staining dishes These dishes are available in a variety of sizes. The small grooved Coplin jars with glass lids are used to stain small number of slides, usually 5-10 slides. Large staining troughs with separate transfer baskets or racks enable up to 20 slides to be stained at the same time.

Staining racks Staining racks are usually made of two pieces of stout glass rods, about 2-4 cm apart and fastened at the ends with rubber tubing or specially designated and adjustable clips. The racks are laid on the bench across the sink. The slides are placed across the rods and stained individually by applying stains and reagents from dropping bottles or pipettes. The racks are ideal for prolonged staining procedures. Staining machines

Staining machines follow the same general pattern of tissue processing machine but instead of tissue containers they carry slide racks. They can be programmed for varying staining periods. These machines have become standard equipments in most histopathology and cytology laboratories 

Staining frozen sections Frozen sections may be stained individually by being transferred from one dish of reagent to the other with a hockey-stick ( a tapered glass rod bent at one end to form an angle of 60°). The sections are attached to the slide at the completion of staining. Sections can also be attached to the slide before staining.

METHODOLOGY OF STAINING

Paraffin sections, after they have been cut, are attached on to the slides and stained. Before staining, certain preparatory treatment of the sections is necessary and involves the following steps: 

1. Removal of paraffin wax (dewaxing) Because paraffin wax is not permeable to stains, it is removed with xylene. Two or three minutes immersion in each of two changes of xylene is usually sufficient for sections up to ten microns thick. This step can be facilitated by first warming the slide in a 60°C oven until the wax just begins to melt. 

2. Removal of xylene Xylene is not miscible with watery solutions and low grade alcohol. It is therefore necessary to remove it with absolute alcohol. One or two minutes in each of the two changes of absolute alcohol is adequate. 

3. Gradual hydration with lower grade alcohol Following treatment of the sections in absolute alcohol, the sections are immersed for a minute or two each in 90% and 70% alcohols. This is to avoid the possibility of diffusion currents bringing damage to, and perhaps detachment of the sections.

4. Hydration with water The sections are now rinsed thoroughly in distilled water or tap water. After this step, the sections are ready to be stained. All the procedures listed above are usually described as 'de-wax and hydrate' or 'section down to water'. In case of stains such as Weigert's elastic stain, it is usual to take the sections from the grade of alcohol nearest to the stain which in the case of Weigert's is absolute alcohol. These steps are common to most staining techniques involving paraffin wax sections. Deviations from the steps will be indicated in the methodologies to be fol lowed. 

5. Removal of artefact pigment Artifact pigments may be present following fixation in formalin or mercuric chloride. They should be removed by the standard methods of saturated picric acid to remove formalin pigment or alcoholic iodine and hypo treatment to remove mercuric chloride artifact.

STAINING IN GENERAL

Staining may be carried out using a single (simple or compound) staining solution or by using two or more separate stains with washing and differentiation in between. Staining time ranges from a few minutes to several hours, starting from dewaxing of the section to mounting of the stained section.

Haematoxylin staining is either progressive or regressive accompanied by washing in tap water or Scotts tap water substitute or lithium carbonate (blueing). If regressive method has been employed, 0.5 % acid alcohol (0.5 ml HCl in 70 % alcohol) is used for differentiation, and this is followed by more washing in water before subsequent staining.

Just as some preparatory steps are taken before performing the actual staining, there are certain steps to be taken after staining before examining the section under the microscope.

Dehydration After staining, paraffin wax sections are usually mounted in a medium that is miscible with xylene or toluene. Since water is not miscible with any of these clearing agents, it becomes necessary to remove all the water (dehydration) through graded alcohols (70%, 90% and two changes of absolute alcohol) before passing the sections into two changes of xylene. Any trace of water in the section when placed in xylene will render the section opaque resulting in the loss of details.

Certain stains are soluble in alcohol, especially the low grade alcohols and for this reason, the sections should not be left in alcohol for more time than is necessary, usually a minute or less is sufficient.

 Clearing in xylene One minute to two in each of two changes of xylene is usually adequate to give transparency to the section and completely remove alcohol from the section. The section should be returned to absolute alcohol if it is found to be insufficiently dehydrated.

MOUNTING STAINED SECTIONS

Due to the great difference in the refractive index of the glass slide, the tissue components and air, an unmounted stained section will show very little details when examined under the microscope. To obtain the best result with stained sections therefore, they have to be mounted in a transparent medium that has a refractive index close to that of the glass slide. Mounting medium is also desired to protect the stained section from physical damage, and from fading of the stain due to heat or oxidation 

Qualities of a good mounting medium are:

1. It must have a refractive index close to that of glass (1.518).

2. It must be freely miscible with xylene and toluene.

3. It must be non-reactive.

4. It will not change the colour or pH

5. It will set hard without granularity or cracking.

6. It will not leak out any stain.

7. It should not cause loss of staining over long periods.

Most mounting media are used for permanent preparation but a few others are temporary or semi-permanent mounting media.

PERMANENT MOUNTING MEDIA

These are usually resins which are either natural or synthetic.

Natural Resins

Canada balsam The most well known natural mounting medium is the canada balsam. It is an oleoresin collected from blisters in the bark of the balsamea (Abies balsamea) fir tree which is native to the eastern half of Canada. It is prepared by dissolving in xylene to form a fairly thin solution (about 40-50%). It is transparent, almost colourless in thin layers, glues firmly to glass and sets to a hard consistency without granulation. Its refractive index is 1.52.

The medium has two disadvantages:

1. It darkens with age and,

2. With time, it slowly becomes acidified. This is because it oxidises xylene to toluic and phthalic acids, and this acidity makes most stains to fade. Because many attempts to neutralise the acidity have failed, the medium is now being superseded by the more stable synthetic resins.

Gum damar It is a natural resin obtained from the tree Sherea wiesveri found in East India. It must be purified before use. It does not change colour with age.

Synthetic Resins

Most of the synthetic mounting media are commercially available. The actual formulation or preparation is kept as a trade secret. But generally, they are prepared by dissolving a polystyrene in an aromatic hydrocarbon solvent to which is added a plasticiser such as dibutylphthalate or tricresy! phosphate to prevent the formation of air spaces when dry

The most common synthetic mountants are: 

1. DPX of Kirkpatrick and Lendrum This is made of 10 g *Distrene 80' (lubricant-free) in 35 ml of xylene and plasticised with 5 ml of dibutyl phthalate. It is a clear colourless mounting medium. DPX shrinks somewhat on drying. Its refractive index is 1.52. 

2. Clarite This is a very popular mounting medium in North America. It is used as a 60% solution in xylene and has a refractive index of 1.544. It does not shrink.

Other synthetic mounting media employed by various workers are HSR (Horleco Synthetic Resin), Clear mount and Material.

The technique for Mounting Sections

Following clearing in xylene, clean a coverslip of the appropriate size and place it on a white blotting paper, preferably on a sheet of Whatman No. I filter paper.

2. With a blunt forcep, pick up the slide carrying the section from the xylene bath, clean the ends first so that the diamond inscribed number is visible and the front of the slide (side on which the section is attached) is identified. Then wipe the back of the slide with a clean, dry, and soft dustfree cloth, and wipe the front to within 2 or 3 mm from the margin of the section.

3. With a small glass rod, place the necessary amount of mounting medium on the section. (The ability to determine the necessary amount of mountant is acquired through experience.)

4. Quickly invert the slide and lower it onto the coverslip, applying gentle pressure. As soon as the medium comes in contact with the coverslip it spreads evenly to the edge of the coverslip and covers the whole area of the section which should still be moist with xylene.

5. Turn the slide over and with a mounted needle, square up the coverslip. Wipe excess medium away with the finger nail covered with a fine cloth dipped in xylene.

6. Finally, the slide is labelled indicating the tissue type or stain used and specimen number.

7. Following this, the sections are ready to be examined. Care should be taken not to move the coverslip. Usually, the slides should be placed on hot plates (50°) or in the wax oven for up to 2hours, or better still, if time permits, at 37°C overnight. This is to harden the mounting medium.

Note

1.Air bubbles are a common problem in mounting sections. This may be due to insufficient xylene left on the section. Air bubbles can also form when the xylene is too much. If the air bubbles are many, it is better to remount, but if one or two, it can be removed by applying gentle pressure on the coverslip with a mounted needle.

2. Coverslips come in various sizes and shapes-round, square or rectangle: 22 x 22, 22 x 30, 22 x 40 and 22 x 50 mm. This variety in sizes and shapes helps to ensure that any given stained section is adequately covered. Occasionally, one may notice cloudy areas in the mounted section. These are caused by moisture. If that is the case, then section has to be re-dehydrated. The coverslip is soaked off in xylene and the slide dehydrated in two baths of absolute alcohol, passed through two baths of xylene and then remounted. The causes of such inadequate dehydration may be due to rapid changes in passage through the alcohols.

Haematoxylin Staining Solutions and Methods