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Tissues from the body taken for diagnosis of disease processes must be processed in the histology laboratory to produce microscopic slides that are viewed under the microscope by pathologists. The techniques for processing the tissues, whether biopsies, larger specimens removed at surgery, or tissues from autopsy, are described below. The persons who do the tissue processing and make the glass microscopic slides are histotechnologists.
Tissue specimens received in the surgical pathology laboratory have a request form that lists the patient information and history along with a description of the site of origin. The specimens are accessioned by giving them a number that will identify each specimen for each patient.
Tissues removed from the body for diagnosis arrive in the Pathology Department and are examined by a pathologist, pathology assistant, or pathology resident. Gross examination consists of describing the specimen and placing all or parts of it into a small plastic cassette which holds the tissue while it is being processed to a paraffin block. Initially, the cassettes are placed into a fixative.
When a malignancy is suspected, then the specimen is often covered with ink in order to mark the margins of the specimen. Different colored inks can be used to identify different areas if needed. When sections are made and processed, the ink will mark the actual margin on the slide.
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:
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. 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.
There are a number of factors that will affect the fixation process:
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 of formalin-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. One way 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 the fixative 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 a concentration 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.
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 sectioned within 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.
Once the tissue has been fixed, it must be processed into a form in which it can be made into thin microscopic sections. The usual way this is done is with paraffin. Tissues embedded in paraffin, which is similar in density to tissue, can be sectioned at anywhere from 3 to 10 microns, usually 6-8 routinely. The technique of getting fixed tissue into paraffin is called tissue processing. The main steps in this process are dehydration and clearing.
Wet fixed tissues (in aqueous solutions) cannot be directly infiltrated with paraffin. First, the water from the tissues must be removed by dehydration. This is usually done with a series of alcohols, say 70% to 95% to 100%. Sometimes the first step is a mixture of formalin and alcohol. Other dehydrants can be used, but have major disadvantages. Acetone is very fast, but a fire hazard, so is safe only for small, hand-processed sets of tissues. Dioxane can be used without clearing, but has toxic fumes.
The next step is called "clearing" and consists of removal of the dehydrant with a substance that will be miscible with the embedding medium (paraffin). The commonest clearing agent is xylene. Toluene works well, and is more tolerant of small amounts of water left in the tissues, but is 3 times more expensive than xylene. Chloroform used to be used, but is a health hazard, and is slow. Methyl salicylate is rarely used because it is expensive, but it smells nice (it is oil of wintergreen).
There are newer clearing agents available for use. Many of them are based on limolene, a volatile oil found in citrus peels. Another uses long chain aliphatic hydrocarbons (Clearite). Although they represent less of a health hazard, they are less forgiving with poorly fixed, dehydrated, or sectioned tissues.
Finally, the tissue is infiltrated with the embedding agent, almost always paraffin. Paraffins can be purchased that differ in melting point, for various hardnesses, depending upon the way the histotechnologist likes them and upon the climate (warm vs. cold). A product called paraplast contains added plasticizers that make the paraffin blocks easier for some technicians to cut. A vacuum can be applied inside the tissue processor to assist penetration of the embedding agent.
The above processes are almost always automated for the large volumes of routine tissues processed. Automation consists of an instrument that moves the tissues around through the various agents on a preset time scale. The "technicon" tissue processor is one of the commonest and most reliable (a mechanical processor with an electric motor that drives gears and cams), though no longer made. Newer processors have computers, not cam wheels, to control them and have sealed reagent wells to which a vacuum and/or heat can be applied.
Tissues that come off the tissue processor are still in the cassettes and must be manually put into the blocks by a technician who must pick the tissues out of the cassette and pour molten paraffin over them. This "embedding" process is very important, because the tissues must be aligned, or oriented, properly in the block of paraffin.
Alternatives to paraffin embedding include various plastics that allow thinner sections. Such plastics include methyl methacrylate, glycol methacrylate, araldite, and epon. Methyl methacrylate is very hard and therefore good for embedding undecalcified bone. Glycol methacrylate has the most widespread use since it is the easiest to work with. Araldite is about the same as methacrylate, but requires a more complex embedding process. Epon is routinely used for electron microscopy where very thin sections are required.
Plastics require special reagents for deydration and clearing that are expensive. For this reason, and because few tissues are plastic embedded, the processing is usually done by hand. A special microtome is required for sectioning these blocks. Small blocks must be made, so the technique lends itself to small biopsies, such as bone marrow or liver.
Once the tissues have been embedded, they must be cut into sections that can be placed on a slide. This is done with a microtome. The microtome is nothing more than a knife with a mechanism for advancing a paraffin block standard distances across it. There are three important necessities for proper sectioning: (1) a very sharp knife, (2) a very sharp knife, and (3) a very sharp knife.
Knives are either of the standard thick metal variety or thin disposable variety (like a disposable razor blade). The former type allows custom sharpening to one's own satisfaction, but is expensive (more than $100 per blade). The latter cost about $1 per blade and are nearly as good. The advantage of the disposable blade becomes apparent when sectioning a block in which is hidden a metal wire or suture.
Plastic blocks (methacrylate, araldite, or epon) are sectioned with glass or diamond knives. A glass knife can section down to about 1 micron. Thin sections for electron microscopy (1/4 micron) are best done with a diamond knife which is very expensive ($2500).
Microtomes have a mechanism for advancing the block across the knife. Usually this distance can be set, for most paraffin embedded tissues at 6 to 8 microns. The more expensive the microtome ($15,000 to $20,000), the better and longer-lasting this mechanism will be.
Sectioning tissues is a real art and takes much skill and practice. Histotechnologists are the artists of the laboratory. It is important to have a properly fixed and embedded block or much artefact can be introduced in the sectioning. Common artefacts include tearing, ripping, "venetian blinds", holes, folding, etc. Once sections are cut, they are floated on a warm water bath that helps remove wrinkles. Then they are picked up on a glass microscopic slide.
The glass slides are then placed in a warm oven for about 15 minutes to help the section adhere to the slide. If this heat might harm such things as antigens for immunostaining, then this step can be bypassed and glue-coated slides used instead to pick up the sections.
At times during performance of surgical procedures, it is necessary to get a rapid diagnosis of a pathologic process. The surgeon may want to know if the margins of his resection for a malignant neoplasm are clear before closing, or an unexpected disease process may be found and require diagnosis to decide what to do next, or it may be necessary to determine if the appropriate tissue has been obtained for further workup of a disease process. This is accomplished through use of a frozen section. The piece(s) of tissue to be studied are snap frozen in a cold liquid or cold environment (-20 to -70 Celsius). Freezing makes the tissue solid enough to section with a microtome.
Frozen sections are performed with an instrument called a cryostat. The cryostat is just a refrigerated box containing a microtome. The temperature inside the cryostat is about -20 to -30 Celsius. The tissue sections are cut and picked up on a glass slide. The sections are then ready for staining.
The embedding process must be reversed in order to get the paraffin wax out of the tissue and allow water soluble dyes to penetrate the sections. Therefore, before any staining can be done, the slides are "deparaffinized" by running them through xylenes (or substitutes) to alcohols to water. There are no stains that can be done on tissues containing paraffin.
The staining process makes use of a variety of dyes that have been chosen for their ability to stain various cellular components of tissue. The routine stain is that of hematoxylin and eosion (H and E). Other stains are referred to as "special stains" because they are employed in specific situations according to the diagnostic need.
Frozen sections are stained by hand, because this is faster for one or a few individual sections. The stain is a "progressive" stain in which the section is left in contact with the stain until the desired tint is achieved.
Hematoxylin is the oxidized product of the logwood tree known as hematein. Since this tree is very rare nowadays, most hematein is of the synthetic variety. In order to use it as a stain it must be "ripened" or oxidized. This can be done naturally by putting the hematein solution on the shelf and waiting several months, or by buying commercially ripened hematoxylin or by putting ripening agents in the hematein solution.
Hematoxylin will not directly stain tissues, but needs a "mordant" or link to the tissues. This is provided by a metal cation such as iron, aluminum, or tungsten. The variety of hematoxylins available for use is based partially on choice of metal ion used. They vary in intensity or hue. Hematoxylin, being a basic dye, has an affinity for the nucleic acids of the cell nucleus.
Hematoxylin stains are either "regressive" or "progressive". With a regressive stain, the slides are left in the solution for a set period of time and then taken back through a solution such as acid-alcohol that removes part of the stain. This method works best for large batches of slides to be stained and is more predictable on a day to day basis. With a progressive stain the slide is dipped in the hematoxylin until the desired intensity of staining is achieved, such as with a frozen section. This is simple for a single slide, but lends itself poorly to batch processing.
Eosin is an acidic dye with an affinity for cytoplasmic components of the cell. There are a variety of eosins that can be synthesized for use, varying in their hue, but they all work about the same. Eosin is much more forgiving than hematoxylin and is less of a problem in the lab. About the only problem you will see is overstaining, especially with decalcified tissues.
The stained section on the slide must be covered with a thin piece plastic or glass to protect the tissue from being scratched, to provide better optical quality for viewing under the microscope, and to preserve the tissue section for years to come. The stained slide must go through the reverse process that it went through from paraffin section to water. The stained slide is taken through a series of alcohol solutions to remove the water, then through clearing agents to a point at which a permanent resinous substance beneath the glass coverslip, or a plastic film, can be placed over the section.
Some tissues contain calcium deposits which are extremely firm and which will not section properly with paraffin embedding owing to the difference in densities between calcium and parffin. Bone specimens are the most likely type here, but other tissues may contain calcified areas as well. This calcium must be removed prior to embedding to allow sectioning. A variety of agents or techniques have been used to decalcify tissue and none of them work perfectly. Mineral acids, organic acids, EDTA, and electrolysis have all been used.
Strong mineral acids such as nitric and hydrochloric acids are used with dense cortical bone because they will remove large quantities of calcium at a rapid rate. Unfortunately, these strong acids also damage cellular morphology, so are not recommended for delicate tissues such as bone marrow.
Organic acids such as acetic and formic acid are better suited to bone marrow, since they are not as harsh. However, they act more slowly on dense cortical bone. Formic acid in a 10% concentration is the best all-around decalcifier. Some commercial solutions are available that combine formic acid with formalin to fix and decalcify tissues at the same time.
EDTA can remove calcium and is not harsh (it is not an acid) but it penetrates tissue poorly and works slowly and is expensive in large amounts.
Electrolysis has been tried in experimental situations where calcium had to be removed with the least tissue damage. It is slow and not suited for routine daily use.
A number of artefacts that appear in stained slides may result from improper fixation, from the type of fixative, from poor dehydration and paraffin infiltration, improper reagents, and poor microtome sectioning.
The presence of a fine black precipitate on the slides, often with no relationship to the tissue (i.e., the precipitate appears adjacent to tissues or within interstices or vessels) suggests formalin-heme pigment has formed. This can be confirmed by polarized light microscopy, because this pigment will polarize a bright white (and the slide will look like many stars in the sky). Formalin-heme pigment is most often seen in very cellular or bloody tissues, or in autopsy tissues, because this pigment forms when the formalin buffer is exhausted and the tissue becomes acidic, promoting the formation of a complex of heme (from red blood cells) and formalin. Tissues such as spleen and lymph node are particularly prone to this artefact. Making thin sections and using enough neutral-buffered formalin (10 to 1 ratio of fixative to tissue) will help. If the fixative solution in which the tissues are sitting is grossly murky brown to red, then place the tissues in new fixative.
The presence of large irregular clumps of black precipitate on slides of tissues fixed in a mercurial fixative such as B-5 suggests that the tissues were not "dezenkerized" prior to staining. These black precipitates will also appear white with polarized light microscopy.
Tissues that are insufficiently dehydrated prior to clearing and infiltration with paraffin wax will be hard to section on the microtome, with tearing artefacts and holes in the sections. Tissue processor cycles should allow sufficient time for dehydration, and final ethanol dehydrant solution should be at 100% concentration. In humid climates, this is difficult to achieve. Covering or sealing the solutions from ambient air will help. Air conditioning (with refrigerants, not with evaporative coolers) will also reduce humidity in the laboratory. Toluene as a clearing agent is more forgiving of poorly dehydrated tissues, but it is more expensive and presents more of a health hazard than other non-xylene clearing agents
Though alcohols such as ethanol make excellent fixatives for cytologic smears, they tend to make tissue sections brittle, resulting in microtome sectioning artefacts with chattering and a "venetian blind" appearance.
Bubbles under the coverslip may form when the mounting media is too thin, and as it dries air is sucked in under the coverslip. Contamination of clearing agents or coverslipping media may also produce a bubbled appearance under the microscope.
"Floaters" are small pieces of tissue that appear on a slide that do not belong there--they have floated in during processing. Floaters may arise from sloppy procedure on the cutting bench-- dirty towels, instruments, or gloves can have tissue that is carried over to the next case. Therefore, it is essential that you do only one specimen at a time and clean thoroughly before opening the container of the next case.
The best way to guard against unrecognized floaters is to always separate like specimens in the numbering sequence. For example, if you have three cases with prostate chips, separate them in accessioning with totally different specimens such as uterus or stomach. That way, if numbers are transposed or labels written wrong or tissue carried over, then you will have an obvious mismatch. Carrying over one prostate to another, or transposing the numbers of identical tissues may never be recognized.
If reusable cassettes are employed, you must be aware that tissue may potentially be carried over and appear as "floaters" even several days later, when the cassette is re-used. The problem arises when, during embedding, not all the tissue is removed from the cassette. Then, in the cleaning process, not all of the wax is removed. Then, the next person using the cassette does not pay attention to the fact that there is tissue already in the cassette and puts his specimen in it. The floater that appears on the slide will look well-preserved--it should, because it was processed to paraffin.
Always be sure that you properly identify the tissue! This means that you make sure that the patient label on the specimen container matches that of the request slip. An accession number is given to the specimen. This number must appear with the tissue at all times. You must never submit a cassette of tissue without a label. You must never submit a cassette of tissue with the wrong label. Mislabelling or unlabelling of tissues is courting disaster.
The lab should be well-ventilated. There are regulations governing formalin and hydrocarbonds such as xylene and toluene. There are limits set by the Occupational Safety and Health Administration (OSHA) that should not be exceeded. These limits have recently been revised to reduced levels.
Every chemical compound used in the laboratory should have a materials safety data sheet on file that specifies the nature, toxicity, and safety precautions to be taken when handling the compound.
The laboratory must have a method for disposal of hazardous wastes. Health care facilities processing tissues often contract this to a waste management company. Tissues that are collected should be stored in formalin and may be disposed by incineration or by putting them through a "tissue grinder" attached to a large sink (similar to a large garbage disposal unit).
Every instrument used in the laboratory should meet electrical safety specifications (be U.L. approved) and have written instructions regarding its use.
Flammable materials may only be stored in approved rooms and only in storage cabinets that are designed for this purpose.
Fire safety procedures are to be posted. Safety equipment including fire extinguishers, fire blankets, and fire alarms should be within easy access. A shower and eyewash should be readily available.
Laboratory accidents must be documented and investigated with incident reports and industrial accident reports.
Specific hazards that you should know about include:
Bouin's solution is made with picric acid. This acid is only sold in the aqueous state. When it dries out, it becomes explosive.
Many reagent kits have sodium azide as a preservative. You are supposed to flush solutions containing sodium azide down the drain with lots of water, or there is a tendency for the azide to form metal azides in the plumbing. These are also explosive.
Benzidine, benzene, anthracene, and napthol containing compounds are carcinogens and should not be used.
Mercury-containing solutions (Zenker's or B-5) should always be discarded into proper containers. Mercury, if poured down a drain, will form amalgams with the metal that build up and cannot be removed.
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