The following points highlight the top four methods of chromosome studies. The methods are: 1. Fixation 2. Stains 3. Pre-Treatment 4. Method of Making Slides Grease-Free.
Chromosome Studies: Method # 1. Fixation:
Fixation may be defined as the selective preservation of morphological organization and chemical composition of tissues for microscopic observation.
The main effects of fixative are — rendering the cell contents insoluble with the coagulation or precipitation of the protein matters; preventing bacterial decomposition; making the tissue suitable for staining; reducing all shrinkage and distortion; and increasing the visibility of the cell contents.
The most dependable fixatives contain one or more substances that precipitate the proteins of the cells or render them insoluble without actual precipitation.
The fixative should be selected for its specific chemical activity so as to preserve the desired structures to the exclusion of other components that might interfere with analysis. The fixative makes the tissue suitable for staining and this is why specific fixatives are used for specific stains.
Acetic acid, ethanol and formaldehyde are most frequently used as cytological fixatives, either alone or in mixture. As all of them have acidic properties, they are used with basic dyes. Acetic acid does not fix cytoplasmic proteins, precipitates nucleoproteins, destroys Golgi and mitochondria swells and softens the tissue.
Ethanol precipitates proteins, dissolves lipids, shrinks and hardens the tissue. Formaldehyde does not precipitate proteins, dissolves lipids, shrinks and hardens the tissue. For electron microscopes, freeze-drying and freeze-etching techniques of fixation are commonly used.
Some Common Cytological Fixatives:
1. Acetic: alcohol (1 : 1) Contains glacial acetic acid 1 part and absolute ethyl alcohol 1 part
2. Acetic: alcohol (1 : 2) Contains glacial acetic acid 1 part and absolute ethyl alcohol 2 part
3. Acetic: alcohol (1 : 3) Contains glacial acetic acid 1 part and absolute ethyl alcohol 3 part
4. Acetic: alcohol (1 : 4) Contains glacial acetic acid 1 part and absolute ethyl alcohol 4 part
5. 70% absolute ethyl alcohol
6. 45% acetic acid
7. Nawaschin’s fixative (Belling’s modification) —
Chromic acid — 5 gms
Glacial acetic acid — 50 ml
Distilled water —320 ml
Formalin — 100 ml
Distilled water — 275 ml
The two solutions should be mixed in 1: 1 ratio just before use.
8. Liwitsky’s fixative —
1% Chromic acid and 10% formalin should be mixed in 1: 1 ratio just before use.
9. Carnoy’s fixative —
1 part glacial acetic acid, 1 part chloroform and 3 parts absolute ethyl alcohol.
10. Bouin’s fixative —
1 part glacial acetic acid, 5 parts 40% formalin and 5 parts picric acid.
Chromosome Studies: Method # 2. Stains:
As we know that most of the cellular components are colorless and so, in order to render them visible under microscope, they must be suitably stained. Generally, staining involves chemical reactions with proteins and nucleic acids.
The selection of a stain depends upon the nature of the material, the type and pH of the fixative used and the chemical reactivity of the stain.
The dyes used for staining cellular components have two kinds of active chemical groups — Chromophoric groups and auxochromic groups. Chromophoric groups impart colour to the dye and auxochromic groups give the dye its ability to attach to the cellular component and to dissolve and dissociate in water.
A dye may be acidic or basic. An acidic dye combines better at low pH and is commonly used for staining the cytoplasm, particularly proteins. The colour is carried by the anion, its net charge is negative and it stains substances with a basic reaction (acidifilic substance). Acid fuchsin, Congo red, methyl blue etc. are acidic dyes.
In basic dye the colour radical is the cation, it has a net positive charge and is used for staining nucleic acids. Basic fuchsin, methyl green, crystal violet etc. are basic dyes. Substances having an affinity for basic dyes are called basophilic. Some dyes are neither acidic nor basic, and both anions and cations have colour properties. These are neutral dyes, e.g. acridine orange.
The chemical basis for staining is provided by the acidic or basic nature of the dye. There is also a physical basis of staining and this is absorption of the dye on the surface of the cellular structures. The use of mordants influence the absorption of the stain. Mordants such as iron alum may be used before staining (pre-mordanting), along with the stain or after staining (post-mordanting).
Common cytological stains such as carmine or orcein etc. require prior fixation of the cells, so only dead cells are observed under microscope. Stains applied to unfixed cells for the detection of specific structures or substances are called vital stains. Examples are Janus green B for mitochondria, methylene blue for dividing cells and neutral red for vacuoles.
Some Common Cytological Stains:
1. 1% Aceto-carmine Stain:
Carmine is a basic dye, reddish purple in colour and obtained from the female scale insect Coccuscacti (class Homoptera) which lives on the cactus Opuntia coccinellifera.
The bodies of the dried females make cochineal and carminic acid is and obtained by extracting cochineal with boiling water, then treating it with lead acetate to produce lead carminate, which is again treated with sulphuric acid. The dye — carmine — is formed by mixing an alum with the carminic acid.
Take 1 gm. of carmine and dissolve it gradually in 100 ml of boiling 45% acetic acid. Heat the solution for about 15 to 20 minutes, carefully keeping it as simmering point. Allow the solution to cool down to room temperature and filter. Use the filtrate.
Alternatively, the heating (5 to 7 hours) can be done in a water-bath after fixing a condenser (a long narrow glass tube) to the conical flask in which the solution is prepared. Sometimes a little extra dye may be added or even a 2% solution may be prepared.
2. 1% Aceto-orcein Stain:
Orcein is a basic dye obtained from the lichen Rocellatinctoria and Lecanoraporella. It is deep purple in colour obtained from the colourless parent compound orcinol 3, 5-di-hydroxytoluene. Take 1 gm. of orcein and dissolve it gradually in 100 ml of boiling 45% acetic acid.
Heat the solution gently for about 7 to 10 minutes, carefully keeping it at simmering point. Allow the solution to cool down to room temperature and then filter. Use the filtrate. Alternatively, the heating (3 to 5 hours) can be done in a water-bath after fixing a condenser to the conical flask in which the solution is prepared.
3. 2% Aceto-orcein Stain:
Take 2 gms orcein and dissolve in 100 ml of 45% acetic acid following the same procedure as above.
4. 1% Crystal Violet Stain:
Take 1 gm. crystal violet and dissolve it in 100 ml of boiling distilled water. Allow the solution to cool down to room temperature and then filter. Use the filtrate.
5. Preparation of Feulgen Stain:
Take 0.5 gm. basic fuchsin and dissolve it in 100 ml of boiling distilled water. Cool the solution to 58°C and filter. Cool the filtrate down to 26°C (using a refrigerator). Add to it 10 ml N.HCl and 0.5 to 1 gm. potassium metabisulphite. Keep the solution in a conical flask wrapped with black paper, seal the cork (preferably rubber cork) with paraffin and put it in a cold, dry place (refrigerator) for 24 hours.
Within this period the solution becomes straw-coloured due to bleaching caused by SO2 liberated by the action of HCI on potassium metabisulphite. The amount of potassium metabisulphite to be added depends upon its freshness. The older the stock, the greater the amount of the substance to be added. If enough SO2 is not liberated then bleaching remains incomplete and the solution does not turn straw- coloured.
If the solution does not turn straw-coloured after 24 hours, add a pinch of charcoal powder to the solution. Keep the solution in a conical flask wrapped with black paper and sealed with paraffin in a refrigerator. A rubber cork should be used to stopper the conical flask. This is done to prevent it from coming in contact with oxygen and light, which cause the colour to return.
The stopper should be sealed with paraffin whenever the flask is opened and the black paper should never be removed. Charcoal powder makes the solution colourless by absorbing the colouring particles.
But if sufficient bleaching is not caused by SO2 then proper staining becomes difficult. The solution thus prepared is leuco fuchsin and is a clear colourless liquid. It is also called Schiff’s reagent. Diamond fuchsin gives the best type of solution. E. Mark of BDH fuchsin also may be used.
Chromosome Studies: Method # 3. Pre-Treatment:
For the determination of chromosome number and morphology, metaphase is the most ideal stage when the chromosomes are shortest and thickest.
But in metaphase the chromosomes remain crowded at the equator of the metaphase spindle and so their individual identity cannot be easily determined. Disorganization of the spindle apparatus is necessary to spread out and flatten the chromosomes, and this is exactly what is done by pre-treatment.
Pre-treatment disorganizes the spindle apparatus, shortens and straightens the chromosome arms and solidifies the cytoplasmic background to some extent. If pressure is now applied on the cell, chromosomes become scattered throughout the cell and their individual identity can be easily recognized.
The pre-treating chemical increases cytoplasmic viscosity bringing it nearer to the viscosity level of spindle substance and thereby the spindle loses its identity. The exact mechanism of spindle disorganization is understood only in case of colchicine which is one of the most important pre-treating agents.
Colchicine is an alkaloid obtained from the roots of Colchicum autumnale (Liliaceae), the crocus plant, or saffron of Kashmir which grows as a common weed in Kashmir having large saffron coloured flowers. Microtubules comprising the spindle fibers are affected by colchicine.
It prevents the assembly of protein into a microtubule; consequently, in presence of colchicine, spindles cannot be formed in cells and mitosis remains arrested at metaphase (C-mitosis or C-metaphase). Prolonged treatment leads to repeated doubling of chromosome number producing polyploid cells.
Pre-treatment is done prior to fixation of the tissue. Although a number of pre-treating agents are in use, it is difficult to generalize the pre-treatment schedule. For every plant species, the ideal pre-treating chemical, the duration of pre-treatment and the optimum temperature of pre-treatment have to be determined by trial and error method.
Some Common Pre-Treating Agents:
0.02 to 1% aqueous solution is used. Pre-treatment is done for 2 to 6 hours at room temperature. As it causes extreme shortening of chromosome, it is preferred for plants with long chromosomes.
It is obtained from the roots of the plant, Aesculus sp. A saturated aqueous solution is used. Pre-treatment is done for 1 to 2 hours at 4°-10°C. It is suitable for plants with short chromosomes.
3. Para-dichlorobenzene (PDB):
A saturated aqueous solution is used. Pre-treatment is usually done for 3 to 4 hours at 10-14°C. It is suitable for plants with long chromosomes. Some other pre-treating chemicals are coumarin, 8-oxyquinoline, α-bromo-naphthal etc. — all are used in aqueous solution of varying concentrations.
Chromosome Studies: Method # 4. Method of Making Slides Grease-Free:
There are several methods of making slides grease-free. The best method is: Prepare a solution of Potassium dichromate (20 gm.) in distilled water (100 ml) and gradually add conc. sulphuric acid (100 ml) to potassium dichromate solution taken in a glass trough.
Keep the slides in this solution for at least 24 hours. Wash the slides in running water, preferably with some detergent powder. Finally, wash them with distilled water.
Secondly, the slides may be kept in 70% alcohol for 7 days and then washed in running water with some detergent powder.
Cytological preparations should be made on grease-free slides. Such preparations are temporary ones. In order to make them permanent, they are to be passed through various grades. If the slides are not grease-free then the material often gets washed out during this process.
Fixation - types of fixatives
The purpose of fixation is to preserve tissues permanently in as life-like a state as possible. Fixation should be carried out as soon as possible after removal of the tissues (in the case of surgical pathology) or soon after death (with autopsy) to prevent autolysis. There is no perfect fixative, though formaldehyde comes the closest. Therefore, a variety of fixatives are available for use, depending on the type of tissue present and features to be demonstrated.
There are five major groups of fixatives, classified according to mechanism of action:
- Oxidizing agents
Aldehydes include formaldehyde (formalin) and glutaraldehyde. Tissue is fixed by cross-linkages formed in the proteins, particularly between lysine residues. This cross-linkage does not harm the structure of proteins greatly, so that antigenicity is not lost. Therefore, formaldehyde is good for immunohistochemical techniques. Formalin penetrates tissue well, but is relatively slow. The standard solution is 10% neutral buffered formalin. A buffer prevents acidity that would promote autolysis and cause precipitation of formol-heme pigment in the tissues.
Glutaraldehyde causes deformation of alpha-helix structure in proteins so is not good for immunohistochemical staining. However, it fixes very quickly so is good for electron microscopy. It penetrates very poorly, but gives best overallcytoplasmic and nuclear detail. The standard solution is a 2% buffered glutaraldehyde
Mercurials fix tissue by an unknown mechanism. They contain mercuricchloride and include such well-known fixatives as B-5 and Zenker's. These fixatives penetrate relatively poorly and cause some tissue hardness, but are fast and give excellent nuclear detail. Their best application is for fixation of hematopoietic and reticuloendothelial tissues. Since they contain mercury, they must be disposed of carefully.
Alcohols, including methyl alcohol (methanol) and ethyl alcohol (ethanol), are protein denaturants and are not used routinely for tissues because they cause too much brittleness and hardness. However, they are very good for cytologic smears because they act quickly and give good nuclear detail. Spray cans of alcohol fixatives are marketed to physicians doing PAP smears, but cheap hairsprays do just as well.
Oxidizing agents include permanganate fixatives (potassium permanganate), dichromate fixatives (potassium dichromate), and osmium tetroxide. They cross-link proteins, but cause extensive denaturation. Some of them have specialized applications, but are used very infrequently.
Picrates include fixatives with picric acid. Foremost among these is Bouin's solution. It has an unknown mechanism of action. It does almost as well as mercurials with nuclear detail but does not cause as much hardness. Picric acid is an explosion hazard in dry form. As a solution, it stains everything it touches yellow, including skin.
Fixation - factors affecting fixation
There are a number of factors that will affect the fixation process:
- Time interval
Fixation is best carried out close to neutral pH, in the range of 6-8. Hypoxia of tissues lowers the pH, so there must be buffering capacity in the fixative to prevent excessive acidity. Acidity favors formation offormalin-heme pigment that appears as black, polarizable deposits in tissue. Common buffers include phosphate, bicarbonate, cacodylate, and veronal. Commercial formalin is buffered with phosphate at a pH of 7.
Penetration of tissues depends upon the diffusability of each individual fixative, which is a constant. Formalin and alcohol penetrate the best, and glutaraldehyde the worst. Mercurials and others are somewhere in between. Oneway to get around this problem is sectioning the tissues thinly (2 to 3 mm). Penetration into a thin section will occur more rapidly than for a thick section.
The volume of fixative is important. There should be a 10:1 ratio of fixative to tissue. Obviously, we often get away with less than this, but may not get ideal fixation. One way to partially solve the problem is to change thefixative at intervals to avoid exhaustion of the fixative. Agitation of the specimen in the fixative will also enhance fixation.
Increasing the temperature, as with all chemical reactions, will increase the speed of fixation, as long as you don't cook the tissue. Hot formalin will fix tissues faster, and this is often the first step on an automated tissue processor.
Concentration of fixative should be adjusted down to the lowest level possible, because you will expend less money for the fixative. Formalin is best at 10%; glutaraldehyde is generally made up at 0.25% to 4%. Too high aconcentration may adversely affect the tissues and produce artefact similar to excessive heat.
Also very important is time interval from of removal of tissues to fixation. The faster you can get the tissue and fix it, the better. Artefact will be introduced by drying, so if tissue is left out, please keep it moist with saline. The longer you wait, the more cellular organelles will be lost and the more nuclear shrinkage and artefactual clumping will occur.
Fixatives - general usage
There are common usages for fixatives in the pathology laboratory based upon the nature of the fixatives, the type of tissue, and the histologic details to be demonstrated.
Formalin is used for all routine surgical pathology and autopsy tissues when an H and E slide is to be produced. Formalin is the most forgiving of all fixatives when conditions are not ideal, and there is no tissue that it will harm significantly. Most clinicians and nurses can understand what formalin is and does and it smells bad enough that they are careful handling it.
Zenker's fixatives are recommended for reticuloendothelial tissues including lymph nodes, spleen, thymus, and bone marrow. Zenker's fixes nuclei very well and gives good detail. However, the mercury deposits must be removed(dezenkerized) before staining or black deposits will result in the sections.
Bouin's solution is sometimes recommended for fixation of testis, GI tract, and endocrine tissue. It does not do a bad job on hematopoietic tissues either, and doesn't require dezenkerizing before staining.
Glutaraldehyde is recommended for fixation of tissues for electron microscopy. The glutaraldehyde must be cold and buffered and not more than 3 months old. The tissue must be as fresh as possible and preferably sectionedwithin the glutaraldehyde at a thickness no more than 1 mm to enhance fixation.
Alcohols, specifically ethanol, are used primarily for cytologic smears. Ethanol (95%) is fast and cheap. Since smears are only a cell or so thick, there is no great problem from shrinkage, and since smears are not sectioned,there is no problem from induced brittleness.
For fixing frozen sections, you can use just about anything--though methanol and ethanol are the best.