Fixation and Decalcification in histopathology

Fixation and Decalcification

FIXATION

Fixation is the preservation after death of the I shape, structure and chemical constituents of tissues and cells. Soon after death, tissues and cells begin to undergo changes leading to their break down and ultimate destruction. Such changes may be due to the action of enzymes normally present in the tissues themselves and so the changes are termed autolytic action (or self destruction). The changes can also be due to external influences such as bacteria which cause decomposition and putrefaction. These bacteria may be those that constitute the normal flora of the body or pathogens that are present due to a disease process.

These changes can be slowed down by low temperatures or prevented by fixation. Adequate and complete fixation is the foundation of all good histological preparations. Faults in fixation cannot be rectified at any later stage and the finished section can only be as good as its initial fixation. The stabilisation of the protein part of the framework of the cell is an important function of fixation. In addition to preserving the tissues and cells, the fixatives (i.e. the fixing fluid or vapour) also makes them resistant to subsequent treatment during processing.

It is essential that tissues are fixed as soon as possible after death or removal from the body. The tissue should never be allowed to dry as it will shrink.

The length of time required for adequate fixation varies according to the size and consistency of the tissue and the fixative employed. If delay in fixation cannot be avoided, the tissue should be kept moist and chilled. The aim of fixation is to prevent the tissues undergoing adverse changes and to preserve the tissues in as life-like a manner as possible. The ideal fixative, in order to fulfil this aim, should meet the following requirements:

1. Prevent autolysis and putrefaction.

2. Should not add to or remove from the tissue constituents.

3. Should not cause any shrinkage or swelling.

4. Should penetrate the tissue and cells rapidly, evenly and deeply.

5. Prevent distortion by any reagents used subsequently

6. Should impart a suitable hardness and texture to allow for easy sectioning.

7. Should render the tissues receptive of stains.

8. Should be non-toxic, non-corrosive and non-inflammable.

9. Should be cheap and easy to prepare.

10. Should be stable.

11. Should allow for long term storage of specimens.

12. Should allow restoration of some natural colours for museum work and photography,

So far in the search for diagnostic perfection, there is no single fixative that fulfils all of the above requirements. Many substances on their own have individual properties that make them fairly good fixatives. These substances are referred to as simple fixatives. So in order to obtain a fixative which will comply as closely as possible with the conditions required for fixation, it becomes necessary to combine several simple fixatives to achieve the desired effect. The resulting solution is referred to as compound fixative. Compound fixative is therefore by definition, a solution of two or more simple fixatives mixed together in order to obtain the combined effect of their individual actions upon the cells and tissue constituents.

Choosing a Fixative

The choice of a fixative is determined by the nature of the investigation required. Several mixtures have become established as routine general fixatives, but most of them have been devised for a specific purpose and are not suitable for routine use.

Fixatives are usually grouped according to their action upon the cell and tissue constituents. Those which preserve the tissue as well as allow the general microscopical study of the tissue structure, and also allow the various layers of tissues and cells to retain their former relationship with each other are called microanatomical fixatives. Those fixatives which are devised for specific actions upon specific areas of the cell are known as cytological fixatives.

This group of fixatives are further sub divided into nuclear and cytoplasmic fixatives. It is obvious from their titles which parts of the cell each acts upon. Practically, those cytological fixatives with a pH of less than 4.6 or those which contain glacial acetic acid are nuclear fixatives and the reverse is true for cytoplasmic fixatives.

Secondary fixation is a technique whereby considerable improvement over the results obtained by the use of 10% formalin alone is achieved. All specimens are collected in 10% formalin and the blocks selected by the pathologist for processing are then treated with a second, more specialised fixative for a shorter length of time. 

Thus the function of formalin for bulk use, distribution outside the laboratory, frozen section work, storage of gross specimens etc. is enhanced and supplemented by the secondary fixatives. The method is however, not without some shortcomings. It complicates the processing schedule and for preparations such as saturated mercuric chloride solution, extreme caution is needed in handling and it is also necessary to remove pigment from the section. This consideration has made the technique less popular.

Whatever method of fixation is used, a large volume of fixative is required. It is recommended that the volume of the fixative should be about 2040 times that of the tissue. In case of large specimens such as whole spleen this is not practicable. Such large specimens should be sliced at intervals of 2 cm or less to allow adequate penetration of the fixative. For the same reasons, hollow specimens like intestines and stomach, should be slit open.

SIMPLE FIXATIVES

Formaldehyde (HCHO) Formaldehyde is a gas which is soluble in water to approximately 40% by weight. It is produced by the oxidation of methanol. This saturated gaseous solution is available commercially as formalin. The term formalin often brings about confusion. For example, a 1:10 dilution of the concentrated solution may be referred to as either 4% formaldehyde or 10% formalin.

Formaldehyde preserves fats and proteins well with no precipitation. On long storage, particularly at low temperature, a white precipitate of paraformaldehyde forms. This can be prevented by storing formalin at room temperature or it can be removed by filtration though it does not adversely affect the fixing property of the reagent. Formalin is acid in reaction due to the presence of formic acid. Though not harmful, the acid can be neutralised by the addition of a small quantity of magnesium carbonate, or 3-4 drops of sodium hydroxide.

Using magnesium carbonate may result in the sudden release of carbon dioxide which can cause an explosion. For this reason a wide mouth vessel should be used for neutralisation and later stored in a Winchester quart bottle. It is a reducing agent and therefore not compatible with potassium dichromate and osmium tetroxide solutions.

Mercuric chloride (HgCl₂) Mercuric chloride is a constituent of many fixatives. It is often used as saturated aqueous solution. It is a good protein precipitant which rapidly penetrates and hardens the tissue. Unfortunately the rate of penetration de creases after the initial few millimeters, as a result large pieces of tissues tend to be hard and overfixed at the periphery and soft and underfixed in the middle. Because of this, and because it causes shrinkage, mercuric chloride is seldom used alone.

Fixatives containing mercuric chloride are listed among the intolerant or harsh fixatives because exposure of tissue to their action in excess of the recommended times will produce excessive hardness and make the cutting of thin sections more difficult. It is radio-opaque and its presence in calcified tissue precludes the use of x-ray to test the end point of decalcification. Its presence in tissues inhibits freezing and makes frozen sections difficult to prepare, even though it neither destroys or preserves lipids. Staining of cytoplasm is en hanced following mercuric chloride fixation; and metachromatic staining is improved and better differentiated.

Tissues fixed in mercuric chloride containing fixative will require treatment to remove the brownish black pigment which always forms. This is easily done by treating the sections with Lugol's iodine for a few minutes and decolourising in sodium thiosulphate prior to staining. Mercuric chloride corrodes most metals and so the fixative should be kept in plastic containers. It is poisonous and should be handled with great care.

Picric acid (C₆H₂ (NO₂)₃OH or trinitrophenol) This fixative is explosive when dry and therefore must be stored damp, preferably under water. It is normally used as saturated aqueous solution at approximately 1% concentration.

Picric acid precipitates proteins and combines with them to form picrates, some of which are soluble in water. For this reason, tissues fixed in picric acid containing fixatives are transferred directly to alcohol to render those picrates insoluble. Glycogen is well preserved. Picric acid causes shrinkage and it is not ideal for fixing kidney due to extreme distortion produced. It imparts yellow colour to tissues so that tiny tissues are easily picked out.

Osmium tetroxide (OsO₄-Osmic acid) It is a pale yellow substance, soluble in water up to 6% at 20°C. The solution is a strong oxidising agent. The aqueous solution is easily reduced by light or by contamination with organic matter. For this reason, it should be stored in a chemically clean bottle in the dark at 2-6°C.

Osmium tetroxide fixes lipids permanently. It preserves cytoplasmic structures well and that is why it is widely used in electron microscopy. It is usually used in combination with chromium salt. It has a poor penetrating power and so it is used to fix small pieces of tissues. Sometimes the vapour of osmium tetroxide is used to fix some tissues such as adrenal because it penetrates better than the solution. Thus washing in running water becomes unnecessary and the production of artefacts is highly reduced. The vapour is however hazardous as exposure to it can result in the deposition of the black oxide (OsO₂) in the cornea and may lead to blindness.

Acetic acid (CH₃ COOH) Acetic acid is usually referred to as glacial acetic acid because at approximately 17°C, it solidifies. It is a colorless solution with a pungent smell. Acetic acid is not used as a fixative alone because of its swelling effect on tissues especially collagen fibers. Its use in compound fixatives is to counteract the shrinking effect of other reagents. Acetic acid is a powerful nucleoprotein precipitant. It destroys mitochondria and golgi element and destroys the lipid-fixing Property of potassium dichromate when used together. It is often used by cytologists to study chromosomes. 

Ethyl alcohol (C₂H₅OH) Ethyl alcohol is colourless, miscible with water and highly inflammable. It is of little use as a simple fixative except for smears and the occasional blood film; but it is frequently incorporated into compound fixatives. It penetrates slowly and tends to harden tissues as well as cause shrinkage which makes staining difficult. Alcohol denatures protein and tends to dissolve lipids. Glycogen is precipitated but not fixed. Being an oxidising agent it should never be used in conjunction with potassium dichromate or osmium tetroxide. 

Potassium dichromate (K₂Cr₂0₇) It is a widely used simple fixative. Depending on the pH of the solution, two different forms of fixatives can be effected. At a pH of less than 4.6, it precipitates protein and fixes carbohydrates. At pH higher than 4.6, the cytoplasm is well preserved and the mitochondria fixed. It is very useful in the study of myelinated nerve fibres. Tissues fixed in a potassium dichromate containing solution should be thoroughly washed in running tap water prior to dehydration to prevent the formation of an insoluble precipitate.

Chromic acid It is a solution of chromium oxide (CrO₃) in water. It is stored as 2% stock solution. It is a powerful oxidising agent and so should not be combined with alcohol or formalin. Tissues fixed in chromic acid should be thoroughly washed in running water to avoid the formation of an insoluble precipitate. 

Acetone Acetone is mainly used in fixatives that allow the demonstration of enzymes. Acetone is similar in its fixing property to alcohol. Glycogen is not well preserved.

COMPOUND FIXATIVES

Broadly, compound fixatives can be treated under two headings:

(1) Micro anatomical and  (2) Cytological fixatives. Micro anatomical fixatives are used to preserve various layers of tissues and cells in relation to one another so that the general structure may be studied. Cytological fixatives are usually sub-divided into nuclear and cytoplasmic fixatives. They are used to preserve the constituent elements of the cells.

MICRO ANATOMICAL FIXATIVES

10% Formal Saline
Sodium chloride	8.5 g
40% formaldehyde	100 ml
Distilled water	900 ml

This is a micro anatomical fixative that is not a compound one. It is probably the most widely used micro anatomical fixative. It is suitable for the fixation of surgical and post-mortem tissues and produces even fixation with very little shrinkage. It is useful for tissues intended as museum specimens as it allows the natural colour to be restored to the specimens. Formal saline is very tolerant and it allows the tissue to be left in it for long periods without excessive hardening or damages, and permits easy sectioning and most staining techniques.

Tissues containing much blood and fixed for a long period in formal saline may contain a fixation artefact, formalin pigment or acid formaldehyde chromatin, which should be removed with picric acid before staining the sections.

Formal saline is the fixative of choice for tissues undergoing subsequent decalcification, for the fixation of brain and other neurological tissues. It is also useful for the fixation of tissues from which frozen sections are to be prepared. Formal saline is a slow fixative and tissues fixed in it may shrink during dehydration but the shrinkage can be reduced by secondary fixation in formal saline sublimate. It is not very suitable for amyloid as the metachromatic reaction is reduced, while some acid dyes stain less intensely than they do following mercuric chloride fixation. Formalin vapour is very irritating and may cause damage to the nasal mucosa or cause sinusitis. It is advisable to wear rubber gloves when handling tissues fixed in formal saline to prevent dermatitis as a result of prolonged contact of formalin with skin. 

Buffered Formalin (or Neutral Buffered Formalin) 

Sodium dihydrogen phosphate (anhydrous)	3.5 g
Disodium hydrogen phosphate (anhydrous)	6.5gm
40% formaldehyde	100 ml
Distilled water	900 ml
Adjust pH to	7.0–7.2

This is a micro anatomical fixative which is specially suitable for the prolonged fixation of routine, surgical, post-mortem and research specimens. Fixation time is 24 hours or longer depending on the size and consistency of the tissue.

The tissue should be washed in running tap water prior to cutting frozen sections or dehydration commencing in 70% alcohol. One big advantage of buffered formalin is the non-formation of acidformaldehyde-haematin pigment. This fixative is also used extensively for tissues which require immunocytochemistry.

Bouin's fluid 

Acetic acid	5 ml
40% formaldehyde	25 ml
Picric acid (saturated)	75 ml

This fixative is used for detailed nuclear studies and the demonstration of glycogen. It is the fixative for embryos. Fixation of small pieces of tissue is complete in 12-24 hours and these tissues should be transferred directly to alcohol without washing in water. This is in order to render the picrates formed insoluble in water. The fixative keeps well, penetrates poorly but rapidly and evenly. It causes little shrinkage. It imparts yellow colour to tissues which is an advantage when dealing with tiny pieces of tissues. Staining of connective tissue by

the trichrome methods is improved but when the - fixative is employed for fibrous tissues, the acetic acid should be left out.

A disadvantage of the fixative is that it should not be used to fix kidney due to the gross distortion it produces. Bouin's fluid is not reliable for cytological observation due to its acetic acid content since structures such as mitochondria may be distorted or totally destroyed.

Zenker's fluid

Sodium sulphate	1.0 g
Potassium dichromate	2.5 g
Mercuric chloride	5.0g
Distilled water	100 ml
Acetic acid	5 ml

This is a general fixative which is highly recommended for small pieces of liver and spleen. It gives excellent cytological detail and brilliant staining of nuclei and connective tissue fibres with the acid aniline dyes. Penetration is poor and tissues should not exceed 0.5 cm in thickness. Fixation is complete in 24 hours. When the tissue is immersed in this fixative for longer than the specified time, it becomes brittle and makes sectioning difficult. Sections must be treated with iodine to remove mercuric artefact pigment.

Helly's fluid (Zenker formol) 

Sodium sulphate	1.0g
Potassium dichromate	2.5g
Mercuric chloride	5.0 g
Distilled water	100 ml
50% formaldehyde	5 ml (add just before use)

Helly's fluid is widely used both as microanatomical and cytological fixative. It is recommended for the fixation of pituitary tissue and bone-marrow. Fixation, though slower than in Zenker's fluid, is complete within 24 hours. Prior to dehydration, the fixed tissue must be thoroughly washed in running water to remove all traces of dichromate. Although the fixative contains an oxidising agent, potassium dichromate and a reducing agent, formaldehyde, it gives excellent nuclear fixation. Staining of the nuclei is more intense than after fixation with Zenker's solution and cytoplasmic granules are well preserved. Tissues left in the fixative for longer than the specified period form a brown scum over the surface. Removal of mercuric pigment with Lugol's iodine must be carried out.

Heidenhain's "Susa" 

Sodium chloride	0.5 g
Mercuric chloride	4.5 g
Trichloracetic acid	2.0 g
Acetic acid	4.0 m
40% formaldehyde	20.0 ml
Distilled water	80.0 ml

This is a good general fixative that is widely used for routine surgical tissues and fibrous tumours. It causes minimal shrinkage and hardening. Fixation of small tissues is usually complete in 1224 hours. After fixation, tissues are transferred directly to 95% or absolute alcohol. Tissues left longer than 24 hours tend to be excessively hardened. Following Susa fixation, many staining procedures including silver impregnation, are permitted giving enhanced results with sharp nuclear details. Fibrous tissues are easy to section. Weigert's elastic fibre stain is, however, difficult to perform satisfactorily. As a routine, sections should be treated for mercuric artefact pigment.

Gendre's solution

Acetic acid, glacial	5.0 ml
Picric acid, saturated solution in 95% alcohol	80.0 ml
Concentrated formaldehyde solution (40%)	15.0 ml

This is a general microanatomical fixative that is also widely used for the preservation of glycogen. It is similar to Bouin's fluid in action but due to the combined action of both alcohol and picric acid, it is superior for the preservation of glycogen. Following fixation, the tissue is washed in several changes of 80% alcohol.

Cytological fixatives These fixatives are divided into two groups: nuclear and cytoplasmic.

Nuclear fixatives 

Carnoy's fluid 

Glacial acetic acid	10.0 ml
Chloroform	30.0 ml
Absolute alcohol	60.0 ml

This nuclear fixative is widely used to fix chromosomes, lymph modes and urgent biopsies. It is rapid in action and also dehydrates thereby allowing for urgent diagnosis to be made. Following fixation, the tissues are transferred directly to absolute alcohol. Fixation period ranges from half an hour to three hours. Glycogen and Nissl granules are well preserved while lipids are destroyed. It causes excessive shrinkage and is only suitable for small biopsies. Red blood corpuscles are haemolysed.

Cytoplasmic fixatives

Flemming's fluid

Flemming's fluid
A Osmium tetroxide	2g
Distilled water	100ml
B Chromium trioxide	1g
Distilled water	100 ml
Working Solution:
Solution A	16 ml
Solution B	60 ml
Acetic acid	4 ml

This perhaps is the most widely used fixative for the preservation of nuclear structures, especially chromosomes. With the omission of acetic acid, the solution becomes a useful cytoplasmic fixative. Small pieces of tissue not more than 2 mm in thickness, are adequately fixed in 12-24 hours. It preserves fat permanently. It is a costly fixative and has a poor penetrative power and so should be used for only small pieces of tissues. It deteriorates rapidly and should be prepared just before use. Following fixation, the tissue should be washed in running tap water for 6-24 hours. Flemming's fluid minus acetic acid is very good for mitochondria and other cytoplasmic structures. Fixation is usually complete within 48 hours. The omission of acetic acid enhances cytoplasmic details. 

Post chromatisation This technique was introduced by the German cytologist, Bendar in 1901 during his research into the nature and structure of mitochondria. This is the treatment of tissues fixed mainly in formal saline with 3% potassium dichromate.

Post chromatisation (also known as post chromining) may be carried out either before processing, when the tissue is left for 5-7 days in the dichromate solution, or after processing when sections, before staining, are immersed in dichromate solution for up to 24 hours, followed in each case by washing well in running tap water.

The purpose is to mordant the tissue. It allows good cytoplasmic staining and gives improved preservation of myelin.

FIXATION OF SMEARS

For the study of exfoliative cytology, smears are fixed in special fixatives designed to minimise distortion of cells.

Alcohol-ether (Equal volumes of 95% alcohol and ether) This is the routinely used cytological fixative for wet smears. It is specially recommended for use with the Papanicolaou staining technique. Smears are fixed within 30 minutes but can be left in the fixative for longer period. Smears are rinsed in water before staining.

Alcohol 95% ethyl or methyl alcohol is an excellent fixative for both wet and dry smears. It is used in the same way as the ether-alcohol mixture.

Schaudinn's Fluid

Schaudinn's Fluid
Mercuric chloride, saturated
aqueous solution	60 ml
Absolute ethyl alcohol	30 ml
Glacial acetic acid	5 ml

This is a rapidly penetrating fixative used for preserving wet smears which are to be stained by haematoxylin and eosin. Smears are well fixed in 2-5 minutes. Following fixation, smears are rinsed in water, treated for mercuric chloride pigment and then stained by the selected method. 

Aerosol spray fixatives Commercial alcohol based fixatives in aerosol spray forms are available. They contain a water soluble wax which provides a protective layer over the smears.

Transport fixative 

Carbowax	3.0g
Glacial acetic acid	0.2 g
Absolute ethyl alcohol	100 ml

This is also called carbowax fixative and it is used to transport smears to the laboratory. The smear is flooded with the fixative and the fluid allowed to evaporate (10-15 minutes). Immerse the smear in absolute ethyl alcohol for at least 10 minutes to remove the carbowax before staining.

FIXATION OF GROSS SPECIMENS

It may be necessary sometimes to fix whole organs. The 10% formal saline fixative or any of the museum fixatives made up of formaldehyde in conjunction with various acetates may be used. For example, 

Sodium acetate	40 g
Formaldehyde 40%	100 ml
Distilled water	900 ml

Fixation techniques depend on the type and size of organ.

Central nervous system After death, autolytic changes rapidly take place in tissues of the central nervous system. For this reason, the tissues are to be fixed as soon as after death as possible. A whole brain is preserved by being suspended in 10% neutral formal saline by means of a thread under the basilar artery. When required whole, the spinal cord should be laid flat on a narrow strip of wood or cork. The dura mater is then incised lengthwise, and pinned onto the board with plastic pins (metal pins will rust and distort the tissue). The pinned spinal cord is then floated with the board uppermost, in 10% neutral formal saline to fix.

Heart To ensure an even and proper penetration, the heart is packed with balls of absorbent cotton wool soaked with 10% formal saline. The organ is then immersed in a large volume of formal saline. 

Liver Liver is best fixed by injecting 10% formal saline into the blood vessels of the liver, using a Robert's bronchogram syringe. Following this, the liver is immersed in large volume of 10% formal saline. In a similar fashion, the kidney and spleen are also fixed.

Lungs From an aspirator, 10% formal saline is run into the main bronchiuntil the linings of the lung appear clearly defined. The aspirator should be placed higher than the specimen. The bronchi should be plugged with absorbent cotton wool and the specimen submerged in large volume of 10% formal saline. 

Intestine The intestine is best fixed in 10% formal saline when it is cut open lengthwise and pinned out on a strip of wood or cork. But by and large the method of fixation will depend on the pathology to be studied. It may be necessary to keep the natural shape, in which case, the intestine is stuffed with absorbent cotton wool and then immersed in the fixative.

SECONDARY FIXATION

Some workers prefer to refix tissues fixed in formal saline for further 4-24 hours. The fixatives usually employed as secondary fixatives are formal sublimate, Zenker formol and Heidenhain's Susa. This procedure is claimed to impart firm texture to the tissue and also improve staining results. 

Post mordanting This technique is employed in the study of the central nervous system when the demonstrations of myelin and phospholipids are required. Following primary fixation in formalin, the tissues are immersed in either potassium dichromate or chromium chloride.

Notes

When preserving tissues for indefinite periods, the following points must be borne in mind:

(a) The fixative may not be suitable for long storage, e.g. formalin becomes acid and affects the tissue. 

(b) The fixative and the tissue may react with each other resulting in the break down of tissue structures. To achieve the desired resuit, a post fixing preservative may be used; examples of such preservatives are 30% solution of diethylene glycol and phenoxethol, the latter possesses bactericidal and fungicidal properties. However, it is always better to block out in paraffin wax or in any other embedding medium; alternatively museum mounting fluid can be used if it is a gross specimen. 

2. Simple fixatives are sometimes referred to by some workers by their action on tissue proteins as either coagulant or non-coagulant fixatives. The most popular ones are:

Coagulant	Non-Coagulant
Ethanol	Formaldehyde
Mercuric chloride	Osmium tetroxide
Chromium trioxide	Potassium dichromate
Picric acid	Acetic acid

DECALCIFICATION

The deposition of calcium salts (calcium phosphate, calcium carbonate and calcium fluorite), usually makes the cutting of fine sections by the usual methods, very difficult. The removal of these calcium salts is known as decalcification. The most suitable method of decalcification depends on the strength, temperature and volume of the decalcifying solution as well as on the size, consistency of the tissue and the type of investigation to be carried out.

Calcium salts occur normally in bones and teeth or in some pathological conditions. There are two types of pathological calcification. The first formis referred to as dystrophic calcification in which the calcium salts are deposited in the cells or surrounding tissues which have been damaged or injured by disease, e.g. tuberculosis or cancerous changes.

The second form of calcification is seen in other lesions and may occur as a result of the pathological processes: it is referred to as metastatic deposition. For example, in hyperthyroidism, there is a change in calcium metabolism causing an increase in blood calcium level.

The principle of decalcification is based on the removal of calcium cations via the anions. The anions are derived from the decalcifying solution which is usually an acid. A good decalcifying agent should remove all calcium without any adverse effect on the cells or tissue fibres and with no impairment of subsequent impregnation or staining.

Apart from acids, buffer solutions of pH 4.4-4.5 and good chelating agents like ethylene diamine tetra-acetic acid, make good agents of decalcification. All acid decalcifying solutions are injurious to the organic ground substance of the bone and other tissues which therefore must be protected by adequate fixation before decalcification is commenced. Fixation is best achieved by selecting suitable blocks from the bone or other calcified tissue, about 2 to 4 mm thick with the aid of a fine hacksaw, or fretsaw and placing them in buffered neutral formalin or in any other non-acid fixative (e.g. Zenker or Helly's fluids) for 15 to 24 hours. A long stay in formalin helps the nucleic acid become resistant to the hydrolytic action of the acids used in decalcification.

Effects of Heat, Agitation and Vacuum

Though heat will accelerate the process of demineralisation, it enhances the destructive action of acids. Heat, also when applied to hasten decalcification, reduces nuclear staining, reduces the effectiveness of trichrome and Van Gieson stains and also affects Periodic acid Schiff (PAS) technique. Heat should therefore not be used in decalcification especially in the warm tropical climate. Agitation of the tissue in the acid solution has little or no accelerating effect on the decalcification process. The same can be said of partial or complete vacuum in decalcification, though it eliminates bubbles.

Preservation of haemosiderin

Haemosiderin is generally removed by acids but more haemosiderins are left over after treatment with formic acid. Preservation of haemosiderin in tissues to be decalcified is carried out as follows: 

1. Fix the bone and marrow in 10% formalin to which 2% yellow ammonium sulphidehas been added. 

2. After fixation, decalcify in 5% formic acid.

General Method of Decalcification

1. By means of a waxed thread, the tissue is suspended in the decalcifying fluid, about 50-100 times its volume. The suspension can also be achieved using a gauze platform supported by a glass rod frame in the jar so that the tissue block rests at, or little below, the middle of the fluid.

2. The fluid should be changed once or twice a day until decalcification is complete. It is important to evaluate the decalcification progress regularly to determine the completion of the process.

3. At the completion of decalcification, the tissue is transferred directly to 70% alcohol and given several changes for about 10 hours.

4. Starting from the last change of 70% alcohol, the tissue is then thoroughly dehydrated.

Tests for Completion of Decalcification

End point of decalcification can be tested in a number of ways. The physical methods of probing the tissue with needle, knife or finger nail are not reliable and they damage the tissue and small pieces of bone may be undetected. The X-ray method is excellent but not always convenient and available and it is not useful when radio-opaque metallic salt such as mercuric chloride has been used in fixation. The chemical method is favoured because it is simple and reliable. The method depends on the detection of dissolved calcium in the decalcifying fluid. 

Method

1. Place 5 ml of the used decalcifying fluid in a clean test tube.

2. Alkalinise by placing a small litmus paper in it, and adding strong ammonia water (Sp. gravity 0.88) drop by drop, shaking after each addition till the litmus paper turns blue.

3. If the solution becomes turbid at this stage, it shows that calcium is present in a large amount and the fluid should be changed.

4. If the solution is clear, carry on with the test. Add 0.5 ml of saturated aqueous ammonium oxalate, mix and leave to stand for 20-30 minutes. Any cloudiness at this stage, indicates the presence of calcium and that calls for a fresh change of fluid.

5. If the solution remains clear, then the end point of decalcification has been reached.

Note

1. It is essential to use distilled water to prepare decalcifying fluid in order to prevent false positive reaction due to the presence of calcium ionsin ordinary tap water. 

2. The test outlined above can be used with almost all methods employing acids except Perenyi's fluid. 

3. After each chemical test, the fluid is removed and the jar thoroughly rinsed with distilled water before new fluid is placed in it.

Decalcifying Solutions and Agents

1. Aqueous formic acid 

Formic acid	5 ml
Distilled water	90 ml
40% formaldehyde (optional)	5 ml

This fluid is ideal for post mortem and research specimens. It permits excellent staining results. It is the most widely used decalcifying fluid. Decalcifying time is 2-7 days. Due to its slow action, it is not suitable for urgent work. At a concentration higher than 8%, the action may become quicker but the resultant cloudiness produced interferes with the chemical test for the end point. X-ray as well as chemical test can be used to test for the end point. 

2. Aqueous nitric acid 

Nitric acid	5 ml
Distilled water	95 ml

This fluid is used mainly for routine purposes. It is rapidly acting and carries very little hydrolysis if the tissue is not allowed to remain immersed in the fluid beyond the required time for decalcification. Due to the formation of nitrous oxide, the fluid usually imparts yellow colouration to the tissue which may affect the staining reaction. The addition of 0.1% urea to the acid will temporarily check the formation of yellow colour.

This acid, like formic acid, can be used with 40% formaldehyde, in which case it becomes ideal for biopsies. Following decalcification and cutting of section, the section should be treated with 1% lithium carbonate overnight.

3. Perenyi's fluid 

Nitric acid, 10% aqueous solution	40 ml
Absolute ethyl alcohol	30 ml
Chromic acid, 0.5% aqueous solution	30 ml

The Perenyi's fluid is a well established decalcifying fluid even though it was originally designed as a fixative. Decalcification time is from 2-10 days. Perenyi's fluid is more often employed as a tissue softener before dehydration. It produces no hardening, and preserves cellular details well, thus making the subsequent staining good. Decalcified tissues need not be washed in water and can be transferred directly to 70% alcohol. The fluid is slow in action. The big disadvantage is that the routine chemical test cannot be used to test for end point of decalcification. This is because a precipitate is formed when ammonia is added to the fluid. For this reason, a modified chemical test is used to test the end point:

1. Place 5 ml of used decalcifying fluid in a chemically clean test tube. 

2. Add a small piece of litmus paper to the fluid. 3. Add ammonium hydroxide drop by drop mixing between drops, until the reaction is alkaline. 

4. Add glacial acetic acid drop by drop until the precipitate dissolves. 

5. Add 0.5 ml of saturated aqueous solution of ammonium oxalate. 

6. The absence of any white precipitate in 20 minutes indicates complete decalcification.

Note

Perenyi's fluid is yellow when freshly prepared but becomes clear violet in no time.

4. Ebner's fluid

5% sodium chloride solution	50 ml
Distilled water	50 ml
Hydrochloric acid	8 ml

Ebner's fluid is very ideal for teeth. It is fairly rapid in action. It hastens dehydration as the decalcified tissue can be transferred directly to 90% alcohol. Nuclear staining is slightly affected.

5. Ion exchange resins

The principle of this method is that calcium ions are removed from solution by the resins which in creases the rate of solubility of calcium from the tissue. The incorporation of ion exchange resins in decalcifying solution adds very little or no improvement to the staining results and the method has no special advantage.

Technique 

A layer of the resin about 13 mm thick, is spread over the bottom of the vessel being used and the specimen is allowed to rest on it. The decalcifying fluid, about 20 times the volume of the resins is added. The chemical test is not used to test the end point, instead the X-ray method is used.

6. Chelating agents

Ethylene diamine tetra acetic acid (EDTA)	5.5 g
10% neutral formalin	100 ml

This solution is very slow in action and is useful where urgency is not needed. Decalcification time is about three weeks and the fluid is changed on every three-day interval, reduced to one day towards the end of decalcification. It is employed for detailed microscopical studies. Its use also minimises the formation of histological artefacts. The subsequent staining results are good.

7. Electrolytic decalcification The principle of decalcification by electrolysis is based on the attraction of the positively charged calcium ions to a negatively charged electrode. The calcified tissue is placed in an electrolyte solution composed of 5% HCI and 5-10% formic acid in equal parts. The temperature is raised during electrolysis and may or may not enhance the rate of decalcification. The rise in temperature may also bring about the charring of the tissue and definitely results in distortion and poor nuclear staining. The method is therefore not recommended. 

8. RDC This fluid known as RDC is obtainable from Bethlehem Instruments Ltd. in the U.K. Its formula is a trade secret. The fluid has been found very valuable in all routine work, and more rapid than conventional fluids. 

9. Calex This is another proprietary fluid whose formula is not made public by the manufacturers. It is considered much better than RDC.

10. Ultrasonic decalcification Ultrasonic waves are produced from an ultrasonic generator which is housed in a metal jacket. This is known as an ultrasonic bath. The fluid which consists of 7.5% acetic acid is placed in the bath and then the bone or tissue placed in it. The technique which was first described by Thorpe, Bellamy and Sharp, is said to be seven times faster than the conventional acid treated controls, and that distortion and interference with staining was minimal. Protagonists of this method attribute the fast rate to the high temperature (up to 50°C) and vibration produced during decalcification. 

Treatment of dense fibrous tissue Dense fibrous tissues though not containing calcium salts, are too tough for sectioning. Blocks of such tissues may be softened using Lendrum's method, i.e. adding 4-6% phenol to the dehydrating alcohols. Commercially produced reagents are also available for softening tissue following embedding in paraffin wax.