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Adequate fixation by an appropriate fixative is central to any histologic preparation. Tissue that is inadequately or inappropriately fixed will lead to difficulties in microtomy, staining, and per-forming ancillary tests. These problems may not be correctable at a later stage.
Unfortunately, there is no ‘‘all-purpose’’ fixa-tive. No single fixative is good for all specimens. It is therefore essential that surgical pathology personnel be familiar with a variety of fixatives and their uses. Although the exact mechanism of action of many fixatives is unknown, fixatives can broadly be classified into four groups based on their mechanism of action. The aldehydes, such as formaldehyde and glutaraldehyde, act by cross-linking proteins, particularly lysine resi-dues. Oxidizing agents, such as osmium tetrox-ide, potassium permanganate, and potassium dichromate, also probably cross-link proteins, al-though their precise mechanism of action is un-known. Acetic acid, methyl alcohol, and ethyl alcohol are all protein-denaturing agents. The fourth and final group of fixatives acts by forming insoluble metallic precipitates, and these agents include mercuric chloride and picric acid. The choice of the appropriate fixative is based on the type of tissue being fixed and on projected needs for ancillary tests such as special stains, immunohistochemistry, in situ hybridization, and electron microscopy. Table 2-1 lists some common fixatives, their basic uses, and their advantages and disadvantages.
Ten percent neutral buffered formalin (4% formaldehyde) is the standard fixative used in most laboratories. Formalin tends to remove water-soluble substances such as glycogen, and it is therefore generally not suitable for the fixa-tion of tissues for electron microscopy. Ten per-cent neutral buffered formalin penetrates and fixes tissues at a rate of approximately 2 to 3 mm/24 h at room temperature.
Glutaraldehyde, a common fixative for electron microscopy, is one of the slowest penetrating fixatives. Tissue for electron microscopy should be cut into 1-mm cubes and immediately placed in refrigerated glutaraldehyde. Glutaraldehyde (4%) must be kept refrigerated before use.
Ethyl alcohol (70% to 100%) is seldom used as a primary fixative. It may be useful in fixing tissue for preserving glycogen and for some histochemi-cal studies, but it has several disadvantages. Ethyl alcohol penetrates tissues very slowly, and because it denatures proteins by abstracting water from the tissue, it can cause excessive hardening, tissue shrinkage, and cell distortion. Alcohol can also dissolve fats and should not be used when lipid studies or stains for myelin are being considered. Carnoy’s is a fixative that combines ethanol, chloroform, and glacial acetic acid. It quickly fixes tissues and it is a good fixative for glycogen, plasma cells, and nucleic acids. Because of its quick action, some labora-tories use Carnoy’s to fix biopsies that require urgent processing.
The mercury-based fixatives (e.g., B5) provide excellent nuclear detail and are useful in evaluat-ing lymphomas. Mercury-based fixatives precipi-tate proteins without firmly binding to them. These fixatives generally must be prepared fresh; once fixed, the tissues require special processing in the histology laboratory (iodine treatment to remove the mercury). Overfixation with B5 can cause excessive hardening of the tissue.
Bouin’s, a picric acid-based fixative, is the fixa-tive of choice for testicular biopsies. Picric acid reacts with basic proteins and forms crystalline picrates with amino acids. Therefore, tissues fixed with picric acid-based fixatives retain little affin-ity for basic dyes, and the picric acid must be recovered from the tissue before staining. Picric acid penetrates tissues well and fixes them rap-idly, but it also causes cells to shrink. Picric acid causes DNA methylation; hence, many poly-merase chain reaction (PCR)-based molecular diagnostic tests cannot be performed on tissues fixed with picric acid.
An appropriate fixation technique is just as im-portant as choosing the correct fixative. Appro-priate fixation requires adequate tissue exposure and a duration of fixation sufficient to allow full penetration of the fixative. For most tissues, a volume of fresh fixative 15 times the volume of tissue is needed to fix the tissue adequately within 12 to 18 hours. The rate of fixation var-ies depending on the type of fixative, the type of tissue, and the thickness of the tissue sections. Adipose tissue (due to its hydrophobic nature) and fibrous tissue (due to its density) may require longer periods of fixation when hydrophilic fixa-tives are employed.
There can be no more important tenet of fixa-tion than to do it early. The process of autolysis begins immediately, and even the best fixative can only arrest, not reverse, this process. Small amounts of tissue may arrive in fixative or saline, whereas larger tissues usually arrive fresh. Large specimens generally do not fix well un-less first prepared. Even then, specimens often require a limited dissection to maximize the surface area exposed to the fixative, thereby en-suring adequate fixation. Tissue with a hollow viscous or lumina should be opened and solid tissue partially serially sectioned at 5- to 10-mm intervals. To maintain proper orientation, these partially sectioned tissues can be pinned onto a wax block and floated in a fixation tank. Paper towels can be inserted between the sections. The towels act as a wick, drawing more fixative to the sections, thereby facilitating rapid fixation. In general, tissue submitted for processing should never exceed a thickness of 4 mm, and tissues comprised of adipose or dense fibrous tissue should be no more than 3 mm. Optimally, you should routinely aim to submit your tissue sec-tions as 2 mm slices. There should be at least a 3-mm space between the cassette and tissue on all sides. Cramming oversized tissue to make it fit into a cassette often results in inferior slide preparation, time consuming reprocessing, and ultimately a delay in diagnosis.
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