PREPARATION OF BIOLOGICAL SPECIMENS
Fixation of tissues is the most crucial step in the preparation of tissue for observation in the transmission electron microscope. Fixation consists of two steps: cessation of normal life functions in the tissue (killing) and stabilization of the structure of the tissue (preservation). The goal of fixation is to preserve structure as faithfully as possible compared to the living state. The three most important parameters to remember about fixation are: (1) keep the time between killing and fixation to a minimum. (2) keep the size of the tissue as small as possible without losing information or destroying the tissue. If a large specimen must be fixed, keep one dimension less than 1 mm, or else nick areas of the specimen that can be discarded so that the fixative can penetrate. Effective penetration of fixative is about 0.5 mm for osmium. (3) keep gross tissue deformation to a minimum by using sharp implements and keeping manipulation of the specimen to the minimum necessary.
Every effort must be made to ensure that the tissue is kept moist in a physiological medium until in fixative. Dissecting in fixative can be used if manipulation of the specimen is time consuming. Ideal size of the specimen . or less (actual size).
If the specimen floats, it must be submerged. In some cases, this may be done by dipping the specimen in a wetting agent briefly before fixation or adding a wetting agent to the fixative solution. If a wetting agent is added to the fixative, then try to transfer the specimen to fresh fixative as soon as possible. Since trapped air is often the cause of tissue floating in the fixative, air can sometimes be removed by partial vacuum. However, this is sometimes damaging to the specimen. The most gentle approach is simply to put a plug of Kimwipe below the surface of the fixative, thereby trapping the specimen beneath the surface of the fixative solution.
Once the specimen is fixed, use the same vial throughout preparation. Simply decant (or pipet) the contents of the vial before each change. Some people prefer to use an aspirator to speed this process. Since tissue lost in the aspirator is irretrievable, beginners would be advised not use the aspirator until they are experienced with handling material.
Fixatives are best used fresh. Assume that all fixatives are at least mildly photoactive and that all are sensitive to oxidation in the environment. For this reason, fixatives are kept in the refrigerator or freezer and warmed to room temperature just before use. Commercially prepared solutions are packed in nitrogen gas.
Glutaraldehyde is normally clear and has a sickly sweet smell. (But don't go out of your way to smell it as it is also an effective fixative as a vapor.) Glutaraldehyde can degrade into glutaric acid which is light yellowish in color. Glutaraldehyde fixation can take place at room temperature or in the refrigerator for 2 hours (for extrememely small material), 6 hours (standard) or more if necessary. Some people hold material in GA for weeks to years, and if it degrades the material, the degradation may not be obvious. (Use of GA as a holding solution should be regarded as a last resort.)
Osmium tetroxide is a straw-colored crystal, which when mixed with water results in a similarly colored solution. It should only be used in a fume hood. Osmium is extremely expensive, so each sample should use only 1 - 2 ml of solution during fixation. It is highly labile at room temperature and all glassware used in its preparation should be acid-washed prior to use to remove organic compounds that will cause its breakdown. Osmium tetroxide in the presence of light, heat, or organic materials will be converted to osmium dioxide, a black compound which is ineffective at fixing materials. Used osmium and black osmium solutions should be discarded in a special waste container in the fume hood. Osmium vapors are extremely effective at fixing mucous membranes and should be regarded as a hazard in the lab. All osmium fixation should be conducted in the cold for up to 2 hours -- no longer.
Dehydration is the chemical removal of water from the specimen. Common dehydrating fluids are ethanol and acetone. The potential problems of dehydration are shrinkage of the specimen, plasmolysis, and removal of soluble components from the specimen. Dehydration must be conducted relatively rapidly in order to prevent excessive extraction of alcohol and acetone-soluble compounds, but slow enough to prevent plasmolysis.
Extraction of specimen components is difficult to control. Low molecular weight carbohydrates are particularly susceptible, since carbohydrates are usually poorly cross-linked if at all following fixation. Proteins tend to be cross-linked by glutaraldehyde during primary fixation and the lipids by osmium tetroxide during secondary fixation. The carbohydrates are essentially unfixed. Linked to the problem of extraction is that of shrinkage. Both problems are most serious at low concentrations in the dehydration series. In general, rapid dehydration is best for these reasons.
By 70% alcohol, the tissue no longer shrinks as much, but does begin to harden. In fact, extended periods of dehydration in the higher concentrations of alcohol may make the tissue quite brittle. If a stopping point is needed, most histologists choose 70% to 100% alcohol as a good place to stop for the evening.
If there is evidence of plasmolysis, perhaps additional dehydration steps (and/or longer changes) may be required. Cell membranes sometimes retain some osmotic activity after short periods of fixation. Longer periods of fixation in glutaraldehyde can reduce osmotic sensitivity as well. (Membranes are essentially insensitive to osmotic changes after 48 hours of fixation in glutaraldehyde.) Poor fixation will aggrevate problems with dehydration.
Dehydration at refrigerator temperatures slows the process down a bit and tends to lend some rigidity to the tissue. It may also reduce plasmolysis slightly. Plants are the most sensitive to poor dehydration, and therefore, refrigerated dehydration is preferred for these tissues.
When changing solutions, make sure that the specimen does not dry out. Therefore, most workers do not pipet the bottom of the vial dry between changes, but leave a little liquid to keep the specimens dry. On the basis of relative volumes, this remaining solution is inconsequential.
To mix alcohol solutions, 95% alcohol is used because using 100% would be prohibitively expensive. Using 95% alcohol to make the dehydrating fluids can be confusing even with the aid of a calculator. Therefore, histologists have developed a short-cut for these calculations: the amount of alcohol needed is calculated as if you were using 100% (for instance, to make about 100 ml of 70% alcohol, you add 70 ml to a graduated cylinder). Then the final volume of the solution is reduced by 5%. (In this example, 25 ml of water is added, making 95 ml of 70% alcohol rather than 100 ml). This is as accurate as adding the 73.68 ml of 95% alcohol which would be needed to make 100 ml of 70% solution, and it is much faster.
Infiltration is the replacement of the dehydrating fluid or transition solvent with plastic resin. In the case of Epon 812 and Spurr's resin, changes of 1/3 resin, 2/3 resin, and 100% resin are used in succession. The steps in infiltration are fewer because less damage is done to the specimen during this stage. The goal of infiltration is simply the complete penetration of resin into the specimen.
Solutions of resin and solvent are prepared immediately before use and added to the specimen vial. Waste resin solutions are discarded in a special container in the fume hood. The resins can be kept in the freezer between uses; however, they must be allowed to warm thoroughly before opening the container since water will condense on cold resin.
Use of slow rotation will aid penetration of the resin into the tissue block. The quality of penetration can be judged only after embedding. In general, a slightly longer period of infiltration is better than poor infiltration.
Embedding involves the final positioning of the tissue specimen in the liquid plastic and its subsequent polymerization. Three types of molds are commonly used in embedding: (1) small capsules, like gelatin capsules or special plastic BEEM capsules; (2) flat embedding molds; (3) flat dishes. Each has its own set of advantages and disadvantages. Embedding is the crucial step in determining the orientation of sectioning. BEEM and gelatin capsules are ideal when the specimen has no polarity, or when the specimen will orient itself by gravity. For particulate samples special BEEM capsules with steep walls and a narrow tip are available. Flat embedding molds are preferred when the specimen must be oriented in a particular position in the liquid plastic. This allows accurate cross and longitudinal sections to be prepared and helps in aligning the specimen. The flat dish is useful when the specimen is large, when orienting the specimen is impractical, or when massive amounts of material are scanned to select a sample to section. The small plastic chip containing the desired specimen must then be sawed out of the block and glued to a dowel or piece of cast plastic for sectioning.
Identification of blocks is easily accomplished and absolutely necessary. Using a pencil or typewriter, write the relevant data on a small sheet of paper or notecard stock to insert into the capsule or mold before embedding. With capsules, the following orientations are useful:
Flat embedding molds require smaller labels since the label cannot be wrapped as easily with these.