Specimen Accessioning

Gross Examination

Fixation - Types of Fixatives

The objective of fixation is to preserve cells and tissue constituents in as close a life-like state as possible and to allow them to undergo further preparative procedures without change. Fixation arrests autolysis and bacterial decomposition and stabilizes the cellular and tissue constituents so that they withstand the subsequent stages of tissue processing. Aside from these requirements for the production of tissue sections, increasing interest in cell constituents and the extensive use of immunohistochemistry to augment histological diagnosis has imposed additional requirements. Fixation should also provide for the preservation of tissue substances and proteins. Fixation is, therefore, the first step and the foundation in a sequence of events that culminates in the final examination of a tissue section.

It is relevant to point out that fixation in itself constitutes a major artefact. The living cell is fluid or in a semi-fluid state, whereas fixation produces coagulation of tissue proteins and constituents, a necessary event to prevent their loss or diffusion during tissue processing; the passage through hypertonic and hypotonic solutions during tissue processing would otherwise disrupt the cells. For example, if fresh unfixed tissues were washed for prolonged periods in running water, severe and irreparable damage and cell lysis would result. In contrast, if the tissues were first fixed in formalin, subsequent immersion in water is generally harmless.

A large variety of fixatives is now available, but no single substance or known combination of substances has the ability to preserve and allow the demonstration of every tissue component. It is for this reason that some fixatives have only special and limited applications, and in other instances, a mixture of two or more reagents is necessary to employ the special properties of each. The selection of an appropriate fixative is based on considerations such as the structures and entities to be demonstrated and the effects of short-term and long-term storage. Each fixative has advantages and disadvantages, some are restrictive while others are multipurpose. The requirements of a large through-put diagnostic laboratory are also quite different from those of a research laboratory with small numbers of specimens for specialised structural analysis and less requirement for urgency.

Over the years, various classifications of fixatives have been proposed, with major divisions according to function as coagulants and non-coagulants, or according to their chemical nature into three general categories which include alcoholic, aldehydic and heavy metal fixatives. A modification of Hopwood's classification is shown below:

1. Aldehydes, such as formaldehyde, glutaraldehyde.
2. Oxidizing agents: metallic ions and complexes, such as osmium tetroxide, chromic acid.
3. Protein-denaturing agents, such as acetic acid, methyl alcohol (methanol), ethyl alcohol
   (ethanol).
4. Unknown mechanism, such as mercuric chloride, picric acid.
5. Combined reagents.
6. Microwaves.
7. Miscellaneous: excluded volume fixation, vapour fixation.

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 immunoperoxidase techniques. Formalin penetrates tissue well, but is relatively slow. The standard solution is 10% neutral buffered formalin (4% formaldehyde). 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 immunoperoxidase staining. However, it fixes very quickly so is good for electron microscopy. It penetrates very poorly, but gives best overall cytoplasmic and nuclear detail. The standard solution is a 2% buffered glutaraldehyde.

Mercurials fix tissue by an unknown mechanism. They contain mercuric chloride 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.

Tissue Processing

Stabilized tissues must be adequately supported before they can be sectioned for microscopic examination. While they may be sectioned following a range of preparatory freezing methods, tissues are more commonly taken through a series of reagents and finally infiltrated and embedded in a stable medium which when hard, provides the necessary support for sectioning by microtomy. This treatment is termed tissue processing. Methods have evolved for a range of embedding media and applications. Pre-eminent amongst these is the paraffin wax method, which is considered to be the most suitable for routine preparation, sectioning, staining and subsequent storage of large numbers of tissue samples. The quality of structural preservation seen in the final stained and mounted section is largely determined by the choice of fixative and embedding medium. During tissue processing, loss of cellular constituents and shrinkage or distortion should be minimal. After fixation, post-fixation and preparatory procedures, the four main stages in the paraffin method are dehydration, clearing, infiltration and embedding.

The first step in processing is dehydration. Water is present in tissues in free and bound (molecular) forms. Tissues are processed to the embedding medium by removing some or all of the free water. During this procedure various cellular components are dissolved by dehydrating fluids. For example, certain lipids are extracted by anhydrous alcohols, and water soluble proteins are dissolved in the lower aqueous alcohols.

In the paraffin wax method, following any necessary post fixation treatment, dehydration from aqueous fixatives is usually initiated in 60%-70% ethanol, progressing through 90%-95% ethanol, then two or three changes of absolute ethanol before proceeding to the clearing stage. Duration of dehydration should be kept to the minimum consistent with the tissues being processed. Tissue blocks 1 mm thick should receive up to 30 minutes in each alcohol, blocks 5 mm thick require up to 90 minutes or longer in each change. Tissues may be held and stored indefinitely in 70% ethanol without harm.

Clearing is the transition step between dehydration and infiltration with the embedding medium. Many dehydrants are immiscible with paraffin wax, and a solvent (transition solvent, ante medium, or clearant) miscible with both the dehydrant and the embedding medium is used to facilitate the transition between dehydration and infiltration steps. Shrinkage occurs when tissues are transferred from the dehydrant to the transition solvent, and from transition solvent to wax. In the final stage shrinkage may result from the extraction of fat by the transition solvent. The term clearing arises because some solvents have high refractive indices (approaching that of dehydrated fixed tissue protein) and, on immersion, anhydrous tissues are rendered transparent or clear. This property is used to ascertain the endpoint and duration of the clearing step. The presence of opaque areas indicates incomplete dehydration.

Xylene clears rapidly and tissues are rendered transparent, facilitating clearing endpoint determination. Concerns over the exposure of personnel to xylene relate mainly to the use of the solvent in coverslipping rather than in processing, and xylene substitutes can be used in these circumstances.

Infiltration is the saturation of tissue cavities and cells by a supporting substance which is generally, but not always, the medium in which they are finally embedded. Tissues are infiltrated by immersion in a substance such as a wax, which is fluid when hot and solid when cold. Alternatively, tissues can be infiltrated with a solution of a substance dissolved in a solvent, for example nitrocellulose in alcohol-ether, which solidifies on evaporation of the solvent to provide a firm mass suitable for sectioning.

Embedding is the process by which tissues are surrounded by a medium such as agar, gelatine, or wax which when solidified will provide sufficient external support during sectioning. Tissues are embedded by placing them in a mould filled with molten embedding medium which is then allowed to solidify. Embedding requirements and procedures are essentially the same for all waxes, and only the technique for paraffin wax is provided here in detail. At the completion of processing, tissues are held in clean paraffin wax which is free of solvent and particulate matter.

Sectioning

Frozen Sections

Hematoxylin and Eosin Staining

The H&E combination was proposed shortly after the discovery of eosin in 1871, although aniline blue was the first counterstain to hematoxylin. Originally, eosin was used alone to color tissues, but its role now is almost exclusively in double and multiple staining procedures.

Hematoxylin cannot stain but must be oxidized to hematein (usually at an acid pH with sodium iodate) which acts as the dye. (Despite this, the staining solution formed is still traditionally referred to as hematoxylin.) However, even at this stage except for a few applications, direct staining is unsuccessful and it is necessary to include a mordant for hematoxylin to stain tissues effectively. The combination of mordant and dye is known as a 'lake', and in the case of hematoxylin-mordant, such lakes are often positively charged, behaving as cationic dyes at low pH. Various metal salts have been used as mordants with hematoxylin, but only those containing aluminum, iron or tungsten are still in common use.

Alum hematoxylin solutions have become the standard, universal means of staining cell nuclei for microscopic examination. Practically every section of normal and diseased tissue will be examined and presumptively identified using an alum hematoxylin to color nuclei. The major disadvantage of alum hematoxylin as a stain is its susceptibility to acids, which limits the range of counterstains that can be used. Alum hematoxylin staining is also influenced by other factors, including the concentration and age of staining solutions as well as the fixation and processing to which the tissue was subject.

The addition of aluminum (at a constant pH), commonly used as a mordant for hematein, has the effect of increasing the absorbance and hence darkening the solutions. In this case hematein is indicated by a band at 430 nm, while a 560 nm peak represents the absorption maximum for the alum hematein complexes.

The most frequently used form of eosin is eosin Y (Cl 45380) which gives yellowish-pink shades (from the Greek eos, dawn) and can be prepared as either an alcoholic (2% solubility) or aqueous (40% solubility) solution. Eosin B (Cl 45400), erythrosin B (Cl 45430) and phloxine B (Cl 45410), along with various similar xanthene dyes, can be used as suitable alternatives.

Eosin Y (M.W. 691.9) is a tetrabrominated derivative of fluorescein with maximum absorption in water between 515 and 518 nm. Commercial preparations may also contain fluorescein and tribromofluorescein in sufficient quantities to influence staining color as the dye becomes more pale with less bromine. The molecule carries one negative charge between pH 3 and 5 and two negative charges above pH 5.

Proteins are generally cationic below pH 6 and will thus bind eosin, probably through the bromine groups. The reaction is influenced by fixation with tissues prepared in Zenker's fluid, in particular, staining strongly. The selectivity and strength of eosin staining can also be enhanced by adding a small amount of glacial acetic to the dye solution.

As a counterstain for hematoxylin, aqueous eosin solutions range between 0.5% and 2% in strength (1% is most common). Alcoholic formulations generally contain less eosin. Correctly applied, eosin should stain various tissue structures shades of pink-especially collagen, cell cytoplasm and erythrocytes. The addition of a very small amount of phloxine can further improve the result.

Hematoxylin And Eosin Staining Of Paraffin Sections

Most standard tissue fixatives are suitable. Cut sections at 3-4 µm.

REAGENTS REQUIRED

1. Ehrlich's hematoxylin:
    hematoxylin (Cl 75290)                       6 g
    absolute alcohol                           300 ml
    distilled water                            300 ml
    glycerol 	                               300 ml
    glacial acetic acid	                        30 ml
    ammonium or potassium aluminum sulphate     30 g
    sodium iodate                              0.9 g

Dissolve the hematoxylin in the alcohol before adding the other ingredients in the order given. Then mix the solution overnight. The addition of sodium iodate artificially ripens the hematoxylin so that it may be used immediately. Alternatively, sodium iodate can be deleted and the mixture ripened by exposure to warmth and sunlight for approximately two months. The naturally ripened form has a longer shelf life.

2. Differentiator

    1% hydrochloric acid in 70% alcohol

3. Alcoholic eosin solution

    95% alcohol                                3.9 l
    eosin Y (CI 45380)                           5 g
    phloxine B (CI 45410)                      0.5 g
    glacial acetic acid                         20 ml

    0.04% aqueous ammonia

METHOD

    1. Dewax and rehydrate sections.
    2. Place in hematoxylin solution for 5-10 minutes.
    3. Wash sections in running water.
    4. Differentiate sections in 1% acid-alcohol and then wash well in water. Repeat if more
       stain needs to be removed. This step requires microscopic control to ensure that only
       nuclei are stained.
    5. Rinse ('blue') in ammonia water (or similar) for 1 minute.
    6. Rinse sections briefly in distilled water.
    7. Counterstain in eosin for 2-5 minutes.
    8. Wash well in water.
    9. Dehydrate in ascending alcohol solutions (50%,70%,80%,95% x 2, 100% x 2).
   10. Clear with xylene (3 - 4 x).
   11. Mount coverslip onto the labeled glass slide with Permount or some other suitable
       organic mounting medium.

RESULTS

Nuclei stain blue; cytoplasm, red blood cells and connective tissue stain shades of pink.

Other Stains

Coverslipping

Decalcification