Jumat, 03 Juni 2011

Preparation of Tissues for Microscopic Examination

The most common procedure used in the study of tissues is the preparation of histological sections that can be studied with the aid of the light microscope. Under the light microscope, tissues are examined via a light beam that is transmitted through the tissue. Because tissues and organs are usually too thick for light to pass through them, they must be sectioned to obtain thin, translucent sections. However, living cells, very thin layers of tissues, or transparent membranes of living animals (eg, the mesentery, the tail of a tadpole, the wall of a hamster's cheek pouch) can be observed directly in the microscope without first being sectioned. It is then possible to study these structures for long periods and under varying physiological or experimental conditions. In most cases, however, tissues must be sliced into thin sections and attached on glass slides before they can be examined. These sections are precisely cut from tissues previously prepared for sectioning using fine cutting instruments called microtomes.
The ideal microscope tissue preparation should be preserved so that the tissue on the slide has the same structure and molecular composition as it had in the body. This is sometimes possible but—as a practical matter—seldom feasible, and artifacts, distortions, and loss of components due to the preparation process are almost always present.

Fixation

If a permanent section is desired, tissues must be fixed. To avoid tissue digestion by enzymes present within the cells (autolysis) or by bacteria and to preserve the structure and molecular composition, pieces of organs should be promptly and adequately treated before or as soon as possible after removal from the animal's body. This treatment—fixation—can be done by chemical or, less frequently, physical methods. In chemical fixation, the tissues are usually immersed in solutions of stabilizing or cross-linking agents called fixatives. Because the fixative needs some time to fully diffuse into the tissues, the tissues are usually cut into small fragments before fixation to facilitate the penetration of the fixative and to guarantee preservation of the tissue. Intravascular perfusion of fixatives can be used. Because the fixative in this case rapidly reaches the tissues through the blood vessels, fixation is greatly improved.
One of the best fixatives for routine light microscopy is a buffered isotonic solution of 4% formaldehyde. The chemistry of the process involved in fixation is complex and not always well understood. Formaldehyde and glutaraldehyde, another widely used fixative, are known to react with the amine groups (NH2) of tissue proteins. In the case of glutaraldehyde, the fixing action is reinforced by virtue of its being a dialdehyde, which can cross-link proteins.
In view of the high resolution afforded by the electron microscope, greater care in fixation is necessary to preserve ultrastructural detail. Toward that end, a double fixation procedure, using a buffered glutaraldehyde solution followed by a second fixation in buffered osmium tetroxide, has become a standard procedure in preparations for ultrastructural studies. The effect of osmium tetroxide is to preserve and stain lipids and proteins.

Embedding
Tissues are usually embedded in a solid medium to facilitate sectioning. To obtain thin sections with the microtome, tissues must be infiltrated after fixation with embedding substances that impart a rigid consistency to the tissue. Embedding materials include paraffin and plastic resins. Paraffin is used routinely for light microscopy; resins are used for both light and electron microscopy.
The process of paraffin embedding, or tissue impregnation, is ordinarily preceded by two main steps: dehydration and clearing. The water is first extracted from the fragments to be embedded by bathing them successively in a graded series of mixtures of ethanol and water (usually from 70% to 100% ethanol). The ethanol is then replaced with a solvent miscible with the embedding medium. In paraffin embedding, the solvent used is usually xylene. As the tissues are infiltrated with the solvent, they generally become transparent (clearing). Once the tissue is impregnated with the solvent, it is placed in melted paraffin in the oven, typically at 58–60°C. The heat causes the solvent to evaporate, and the spaces within the tissues become filled with paraffin. The tissue together with its impregnating paraffin hardens after being taken out of the oven. Tissues to be embedded with plastic resin are also dehydrated in ethanol and—depending on the kind of resin used—subsequently infiltrated with plastic solvents. The ethanol or the solvents are later replaced by plastic solutions that are hardened by means of cross-linking polymerizers. Plastic embedding prevents the shrinking caused by the high temperatures needed for paraffin embedding and gives much better results.
The hard blocks containing the tissues are then taken to a microtome (Figure 1–1) and are sectioned by the microtome's steel or glass blade to a thickness of 1–10 m. Remember that 1 micrometer (1 m) = 0.001 mm = 10–6 m; 1 nanometer (1 nm) = 0.001 m = 10–6 mm = 10–9 m. The sections are floated on water and transferred to glass slides to be stained.

Figure 1–1
Microtome for sectioning resin- and paraffin-embedded tissues for light microscopy. Rotation of the drive wheel moves the tissue-block holder up and down. Each turn of the drive wheel advances the specimen holder a controlled distance, generally between 1 and 10 m. After each forward move, the tissue block passes over the knife edge, which cuts the sections. (Courtesy of Microm.)
A completely different way to prepare tissue sections is to submit the tissues to rapid freezing. In this process, the tissues are fixed by freezing (physically, not chemically) and at the same time become hard and thus ready to be sectioned. A freezing microtome—the cryostat (Gr. kryos, cold, + statos, standing)—has been devised to section the frozen tissues. Because this method allows stained sections to be prepared rapidly (within a few minutes), it is routinely used in hospitals to study specimens during surgical procedures. Freezing of tissues is also effective in the histochemical study of very sensitive enzymes or small molecules, since freezing does not inactivate most enzymes. Because immersion of tissues in solvents such as xylene dissolves the tissue lipids, the use of frozen sections is advised when these compounds are to be studied.

Staining
To be studied microscopically most sections must be stained. With few exceptions, most tissues are colorless, so observing them unstained in the light microscope is useless. Methods of staining tissues have therefore been devised that not only make the various tissue components conspicuous but also permit distinctions to be made between them. The dyes stain tissue components more or less selectively. Most of these dyes behave like acidic or basic compounds and have a tendency to form electrostatic (salt) linkages with ionizable radicals of the tissues. Tissue components that stain more readily with basic dyes are termed basophilic (Gr. basis, base, + phileo, to love); those with an affinity for acid dyes are termed acidophilic.
Examples of basic dyes are toluidine blue and methylene blue. Hematoxylin behaves like a basic dye, that is, it stains the basophilic tissue components. The main tissue components that ionize and react with basic dyes do so because of acids in their composition (nucleic acids, glycosaminoglycans, and acid glycoproteins). Acid dyes (eg, orange G, eosin, acid fuchsin) stain the acidophilic components of tissues such as mitochondria, secretory granules, and collagen.
Of all dyes, the combination of hematoxylin and eosin (H&E) is the most commonly used. Hematoxylin stains the cell nucleus and other acidic structures (such as RNA-rich portions of the cytoplasm and the matrix of hyaline cartilage) blue. In contrast, eosin stains the cytoplasm and collagen pink. Many other dyes, such as the trichromes (eg, Mallory's stain, Masson's stain), are used in different histological procedures. The trichromes, in addition to showing the nuclei and cytoplasm very well, help to differentiate collagen from smooth muscle. A good technique for differentiating collagen is the use of picrosirius, especially when associated with polarized light (see Polarizing Microscopy).
In many procedures (see Immunocytochemistry), the sections become labeled by a precipitate, but cells and cell limits are often not visible. In this case a counterstain, usually a single stain that is applied to a section to allow the recognition of nuclei or cytoplasm, is used.
Although most stains are useful in visualizing the various tissue components, they usually provide no insight into the chemical nature of the tissue being studied. In addition to tissue staining with dyes, impregnation with metals such as silver and gold is a common method, especially in studies of the nervous system.
The whole procedure, from fixation to observing a tissue in a light microscope, may take from 12 h to 21⁄2 days, depending on the size of the tissue, the fixative, and the embedding medium.

Light Microscopy
Conventional light, phase-contrast, differential interference, polarizing, confocal, and fluorescence microscopy are all based on the interaction of light and tissue components. With the light microscope, stained preparations are usually examined by means of light that passes through the specimen. The microscope is composed of mechanical and optical parts (Figure 1–2). The optical components consist of three systems of lenses: condenser, objective, and eyepiece. The condenser collects and focuses light, producing a cone of light that illuminates the object to be observed. The objective lenses enlarge and project the illuminated image of the object in the direction of the eyepiece. The eyepiece further magnifies this image and projects it onto the viewer's retina, a photographic plate, or (to obtain a digital image) a detector such as a charged coupled device camera. The total magnification is obtained by multiplying the magnifying power of the objective and eyepiece.

Figure 1–2

Schematic drawing of a light microscope showing its main components and the pathway of light from the substage lamp to the eye of the observer. (Courtesy of Carl Zeiss Co.)

Resolution
The critical factor in obtaining a crisp, detailed image with the microscope is its resolving power, that is, the smallest distance between two particles at which they can be seen as separate objects. The maximal resolving power of the light microscope is approximately 0.2 m; this power permits good images magnified 1000–1500 times. Objects smaller than 0.2 m (such as a membrane or a filament of actin) cannot be distinguished with this instrument. Likewise, two objects, such as two mitochondria or two lysosomes, will be seen as only one object if they are separated by less than 0.2 m. The quality of the image—its clarity and richness of detail—depends on the microscope's resolving power. The magnification is of value only when accompanied by high resolution. The resolving power of a microscope depends mainly on the quality of its objective lens. The eyepiece lens enlarges only the image obtained by the objective; it does not improve resolution. For this reason, when comparing objectives of different magnifications, those that provide higher magnification also have higher resolving power.
Highly sensitive video cameras enhance the power of the light microscope and allow the capture of digitized images that can be fed into computers for quantitative image analysis and printing.
The frontiers of light microscopy have been redefined by the use of video cameras highly sensitive to light. With cameras and image-enhancement programs, objects that may not be visible when viewed directly through the eyepiece may be made visible in the video screen. These video systems are also useful for studying living cells for long periods of time, because they use low-intensity light and thus avoid the cellular damage that can result from intense illumination.
The electronic images from video cameras can be easily digitized and adapted to the specific requirements of an experiment through computer programming. For example, contrast enhancement is an important computer-assisted technique that may reveal to the investigator a structural image not immediately seen when the specimen is observed directly in the microscope. Software developed for image analysis allows the measurement of microscopic structures.

References
Alberts B et al: Molecular Biology of the Cell, 3rd ed. Garland, 1994.
Bancroft JD, Stevens A: Theory and Practice of Histological Techniques, 2nd ed. Churchill Livingstone, 1990.
Cuello ACC: Immunocytochemistry. Wiley, 1983.
Darnell J, Lodish H, Baltimore D: Molecular Cell Biology, 2nd ed. Scientific American Books, 1990.
Hayat MA: Stains and Cytochemical Methods. Plenum, 1993.
James J: Light Microscopic Techniques in Biology and Medicine. Martinus Nijhoff, 1976.
Junqueira LCU et al: Differential staining of collagen types I, II and III by Sirius Red and polarization microscopy. Arch Histol Jpn 1978;41:267. [PMID: 82432]
Meek GA: Practical Electron Microscopy for Biologists. Wiley, 1976.
Pease AGE: Histochemistry: Theoretical and Applied, 4th ed. Churchill Livingstone, 1980.
Rochow TG, Tucker PA: Introduction to Microscopy by Means of Light, Electrons, X Rays, or Acoustics. Plenum, 1994.
Rogers AW: Techniques of Autoradiography, 3rd ed. Elsevier, 1979.
Rubbi CP: Light Microscopy. Essential Data. Wiley, 1994.
Spencer M: Fundamentals of Light Microscopy. Cambridge University Press, 1982.
Stoward PJ, Polak JM (editors): Histochemistry: The Widening Horizons of Its Applications in Biological Sciences. Wiley, 1981.

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