6 Tissue processing
After the removal of a tissue sample from the patient, a series of physical and chemical processes must take place to ensure that the final microscopic slides produced are of a diagnostic quality. Tissues are exposed to a series of reagents that fix, dehydrate, clear, and infiltrate the tissue. The tissue is finally embedded in a medium that provides support for microtomy. The quality of the structural preservation of tissue components is determined by the choice of exposure times to the reagents during processing. Every step in tissue processing is important; from selection of the sample, determining the appropriate protocols and reagents to use, to staining and final diagnosis. Producing quality slides for diagnosis requires skills that are developed through continued practice and experience. As new technology and instrumentation develops, the role of the histology laboratory in patient care will continue to evolve, providing standardization of processes, increased productivity, and better utilization of the resources available. This chapter will provide an overview of the steps in the process and the reagents needed to prepare tissue for microscopic evaluation.
A unique accession number or code should be assigned to every tissue sample as discussed in Chapter 5. This unique number should accompany the specimens throughout the entire laboratory process and may be electronically or manually generated. New technology has made bar code quick response (QR) and character recognition systems readily available in most laboratories. Automated pre-labeling systems that permanently etch or emboss tissue cassettes and slides, as well as chemically resistant pens, pencils, slides and labels, are routinely used in pathology laboratories. Regardless of whether an automated or manual labeling system is used, adequate policies and procedures must be in place to ensure positive identification of the tissue blocks and slides during processing, diagnosis, and filing.
Tissue processing is designed to remove all extractable water from the tissue, replacing it with a support medium that provides sufficient rigidity to enable sectioning of the tissue without parenchymal damage or distortion.
When tissue is immersed in fluid, an interchange occurs between the fluid within the tissue and the surrounding fluid. The rate of fluid exchange is dependent upon the exposed surface of the tissue that is in contact with the processing reagents. Several factors influence the rate at which the interchange occurs: namely, agitation, heat, viscosity and vacuum.
Agitation increases the flow of fresh solutions around the tissue. Automated processors incorporate vertical or rotary oscillation, or pressurized removal and replacement of fluids at timed intervals as the mechanism for agitation. Efficient agitation may reduce the overall processing time by up to 30%.
Heat increases the rate of penetration and fluid exchange. Heat must be used sparingly to reduce the possibility of shrinkage, hardening or embrittlement of the tissue sample. Temperatures limited to 45°C can be used, but higher temperatures may be deleterious to subsequent immunohistochemistry.
Viscosity is the property of resistance to the flow of a fluid. The smaller size of the molecules in the solution, the faster the rate of fluid penetration (low viscosity). Conversely, if the molecule size is larger, the rate of exchange is slower (high viscosity). Most of the solutions used in processing, dehydration and clearing, have similar viscosities, with the exception of cedar wood oil. Embedding mediums have varying viscosities. Paraffin has a lower viscosity in the fluid (melted) state, enhancing the rapidity of the impregnation.
Using pressure to increase the rate of infiltration decreases the time necessary to complete each step in the processing of tissue samples. Vacuum will remove reagents from the tissue, but only if they are more volatile than the reagent being replaced. Vacuum used on the automated processor should not exceed 50.79 kPa to prevent damage and deterioration to the tissue. Vacuum can also aid in the removal of trapped air in porous tissue. Impregnation time for dense, fatty tissue can be greatly reduced with the addition of vacuum during processing.
Preserving cells and tissue components with minimal distortion is the most important aim of processing tissue samples. Fixation stabilizes proteins, rendering the cell and its components resistant to further autolysis by inactivating lysosomal enzymes. It also changes the tissues’ receptiveness to further processing. Fixation must finish before subsequent steps in the processing schedule are initiated.
If fixation is not complete prior to processing, stations should be designated on the processor for this purpose. If the tissue is inadequately fixed, the subsequent dehydration solutions may complete the process, possibly altering the staining characteristics of the tissue. The size and type of specimen in the tissue cassette determines the time needed for complete fixation and processing. The tissue should be dissected to 2–4 mm in thickness. Care must be taken not to overfill the cassette, as this would impede the flow of reagents around the tissue. If possible, larger and smaller pieces of tissue should be separated and processed using different schedules. The most commonly used reagent for the fixation of histological specimens is 10% neutral buffered formalin (NBF) – see Chapter 4.
Special fixation techniques may require additional steps before processing is initiated. Picric acid fixatives (Bouin’s) form water-soluble picrates making it necessary to place the tissue cassettes directly into 70% alcohol for processing. Alcoholic fixatives, such as Carnoy’s fluid, should be placed directly into 100% alcohol. To help in the visualization of small fragments of tissue during embedding, a few drops of 1% eosin can be added to the specimen container 30 minutes prior to processing. The pink color of the tissue remains during processing, but washes out during subsequent staining.
The first stage of processing is the removal of ‘free’ unbound water and aqueous fixatives from the tissue components. Many dehydrating reagents are hydrophilic (‘water loving’), possessing strong polar groups that interact with the water molecules in the tissue by hydrogen bonding. Other reagents affect dehydration by repeated dilution of the aqueous tissue fluids. Dehydration should be accomplished slowly. If the concentration gradient is excessive, diffusion currents across the cell membranes may increase the possibility of cell distortion. For this reason, specimens are processed through a graded series of reagents of increasing concentration. Excessive dehydration may cause the tissue to become hard, brittle and shrunken. Incomplete dehydration will impair the penetration of the clearing reagents into the tissue, leaving the specimen soft and non-receptive to infiltration. There are numerous dehydrating agent; ethanol, ethanol acetone, methanol, isopropyl, glycol and denatured alcohols.
Ethanol is a clear, colorless, flammable liquid. It is hydrophilic, miscible with water and other organic solvents, fast-acting and reliable. Aside from its human health-risk potential, ethanol is taxable, controlled by many governments, and therefore requires careful record keeping. Graded concentrations of ethanol are used for dehydration; the tissue is immersed in 70% ethanol in water, followed by 95% and 100% solutions. Ethanol ensures total dehydration, making it the reagent of choice for the processing of electron microscopy specimens. For delicate tissue it is recommended that the processing starts in 30% ethanol.
This fluid has the same physical property as ethanol. Denatured alcohol consists of ethanol, with the addition of methanol (about 1%), isopropyl alcohol or a combination of alcohols. For purposes of tissue processing it is used in the same manner as ethanol.
Acetone is a clear, colorless, flammable fluid that is miscible with water, ethanol and most organic solvents. It is rapid in action, but has poor penetration and causes brittleness in tissues if its use is prolonged. Acetone removes lipids from tissue during processing.
When added to dehydrating agents, phenol acts as a softening agent for hard tissues such as tendon, nail, and dense fibrous tissue and keratin masses. Phenol (4%) should be added to each of the 95% ethanol stations. Alternatively, hard tissue can be immersed in a glycerol/alcohol mixture.
Universal solvents are no longer used for routine processing due to their hazardous properties, and they should be handled with extreme care. Universal solvents both dehydrate and clear tissues during tissue processing. Dioxane, tertiary butanol and tetrahydrofuran are considered to be universal solvents. They are not recommended for processing delicate tissues due to their hardening properties.
Clearing reagents act as an intermediary between the dehydration and infiltration solutions. They should be miscible with both solutions. Most clearants are hydrocarbons with refractive indices similar to protein. When the dehydrating agent has been entirely replaced by most of these solvents the tissue has a translucent appearance: hence the term ‘clearing agent’.
Most clearing agents are flammable liquids, which warrant caution in their use. The boiling point of the clearing agent gives an indication of its speed of replacement by melted paraffin wax. Fluids with a low boiling point are generally more readily replaced. Viscosity influences the speed of penetration of the clearing agent. Prolonged exposure to most clearing agents causes the tissue to become brittle. The time in the clearing agent should be closely monitored to ensure that dense tissue blocks are sufficiently cleared and smaller more fragile tissue blocks are not damaged. Cost should be considered, especially as it relates to disposal of the reagent. Since most clearing agents are aromatic hydrocarbons or short-chain aliphatic hydrocarbons, environmental issues must be addressed. Most institutions have a policy for the storage, disposal and safety requirements for all flammables used in the laboratory.
Xylene is a flammable, colorless liquid with a characteristic petroleum or aromatic odor, which is miscible with most organic solvents and paraffin wax. It is suitable for clearing blocks that are less than 5 mm in thickness and rapidly replaces alcohol from the tissue. Overexposure to xylene during processing can cause hardening of tissues. It is most commonly used in routine histology laboratories and is also recyclable.
Chloroform is slower in action than xylene but causes less brittleness. Thicker tissue blocks can be processed, greater than 1 mm in thickness. Tissues placed in chloroform do not become translucent. It is non-flammable but highly toxic, and produces highly toxic phosgene gas when heated. It is most commonly used when processing specimens of the central nervous system.
Xylene substitutes are aliphatic hydrocarbons that exist in long- and short-chained forms. They differ in the number of carbon atoms within the carbon chain. Short-chained aliphatics have the same evaporation properties as xylene, and have no affinity for water. Long-chained aliphatics do not evaporate rapidly and may cause contamination of the paraffin wax on tissue processors.
Limonene reagents are extracts from orange and lemon rinds; they are non-toxic and miscible with water. Disposal is dependent upon the water treatment centers and local/national standards. The main disadvantages are that they can cause sensitization and have a strong pungent odor that may cause headaches. Also, small mineral deposits such as copper or calcium may dissolve and leach from tissues. They are extremely oily and cannot be recycled.
Paraffin wax continues to be the most popular infiltration and embedding medium in histopathology laboratories. Paraffin wax is a mixture of long-chained hydrocarbons produced in the cracking of mineral oil. Its properties are varied depending on the melting point used, ranging from 47 to 64°C. Paraffin wax permeates the tissue in liquid form and solidifies rapidly when cooled. The tissue is impregnated with the medium, forming a matrix and preventing distortion of the tissue structure during microtomy. It has a wide range of melting points, which is important for use in the different climatic regions of the world. To promote desirable ribboning during microtomy, paraffin wax of suitable hardness at room temperature should be chosen. Heating the paraffin wax to a high temperature alters the properties of the wax. Higher melting point paraffin wax provides better support for harder tissues, e.g. bone, can allow production of thinner sections, but may cause difficulty with ribboning. Lower melting point paraffin wax is softer and provides less support for harder tissues. It is more difficult to obtain thinner sections but ribboning is easier. Paraffin wax is inexpensive, provides quality sections and is easily adaptable to a variety of uses. Paraffin wax is compatible with most routine and special stains, as well as immunohistochemistry protocols.
Paraffin waxes that contain plasticizers or other resin additives are commercially available, providing a selection that is appropriate for most laboratories. These additives create paraffin waxes with selectable hardness compatible with the tissue to be embedded. The amount of additive will impact the rate of infiltration. Substances added to paraffin wax in the past include beeswax, rubber, ceresin, plastic polymers and diethylene glycol distearate. Many of these additives had a higher melting point than paraffin wax, consequently making the tissue more brittle.
Resin is used exclusively as the embedding medium for electron microscopy (see Chapter 22), ultra-thin sectioning for high resolution and also for undecalcified bone (see Chapter 16).
Agar gel alone does not provide sufficient support for sectioning tissues. Its main use is as a cohesive agent for small friable pieces of tissue after fixation, a process known as double embedding. Fragments of tissue are embedded in melted agar, allowed to solidify and trimmed for routine processing. A superior, more refined, method is to filter the fixative containing small, friable tissue fragments through a Millipore filter using suction. Molten agar is then carefully poured into the filter apparatus, the agar is left to solidify and the resultant agar pellet is removed and routinely processed and embedded in paraffin wax.
Embedding involves the enclosing of properly processed, correctly oriented specimens in a support medium that provides external support during microtomy. The embedding media must fill the matrix within the tissue, supporting cellular components. The medium should provide elasticity, resisting section distortion while facilitating sectioning.
Most laboratories use modular embedding centers, consisting of a paraffin dispenser, a cold plate, and a heated storage area for molds and tissue cassettes. Paraffin wax is dispensed automatically from a nozzle into a suitably sized mold. The tissue is oriented in the mold; a cassette is attached, producing a flat block face with parallel sides. The mold is placed on a small cooling area to allow the paraffin wax to solidify. The quick cooling of the wax ensures a small crystalline structure, producing fewer artifacts when sectioning the tissue.
Specimen orientation during embedding is important for the demonstration of proper morphology. Incorrect orientation may result in diagnostic tissue elements being damaged during microscopy or not being evident for pathology review. Products are available that help ensure proper orientation: marking systems, tattoo dyes, biopsy bags, sponges, and papers. Orientation of the tissue should offer the least resistance of the tissue against the knife during sectioning. A margin of embedding medium around the tissue assures support of the tissue.
• Intestine, gallbladder, and other epithelial biopsies: cut in a plane at right angles to the surface, and oriented so the epithelial surface is cut last, minimizing compression and distortion of the epithelial layer.