Introduction to microtechnique

1.1. Thickness and contrast 1
1.2. Staining and histochemistry 3
1.3. Some physical considerations 4
1.4. Properties of tissues 5
1.5. Books and journals 6
1.6. On carrying out instructions 6
1.6.1. De-waxing and hydration of paraffin sections 6
1.6.1.1. Solvent method 6
1.6.1.2. Emulsification method 7
1.6.2. Staining 8
1.6.3. Washing and rinsing 8
1.6.4. Dehydration and clearing 9
1.6.5. Staining through paraffin: an irrational method 9
1.6.6. Mechanization 10
1.7. Whole mounts and free cells 10
1.8. Understanding the methods 10

Many theoretical explanations and practical instructions are contained in this book. The present chapter concerns aspects of the making of microscopical preparations that are fundamental to all the techniques described in the later chapters. It cannot be over-emphasized that unless the student or technician understands the rationale of all that is to be done, he will not do it properly. Chapter 4 and Section 1.6 of this chapter contain some of the practical information relevant to the manipulations discussed in all parts of the book.

With an ordinary light microscope (LM) it is possible to see only limited structural detail in a living or freshly removed part of a large organism. For the resolution of finer structure within and around cells it is necessary to study fixed specimens. These are pieces of animal or plant material that have been structurally stabilized, usually by a chemical treatment. Fixation, which is reviewed in Chapter 2, arrests post-mortem decay and also gives a harder consistency to many tissues. Fixation introduces structural and chemical artifacts, but these are fairly well understood and for most purposes they outweigh the technical difficulties and artifacts encountered in the examination of unfixed specimens. Some naturally hard materials require softening treatments after fixation (Chapter 3); bone, for example, can be decalcified.

1.1. Thickness and contrast

In order to be examined with a microscope, a specimen must be thin enough to be transparent and must possess sufficient contrast to permit the resolution of structural detail. Thinness may be an intrinsic property of the object to be examined. Thus, small animals and plants, films or smears of cells, tissue cultures, macerated or teased tissues, and spread-out sheets of epithelium or connective tissue are all thin enough to mount on slides directly. In histology, histopathology and histochemistry, one is more often concerned with the internal structure of larger, solid specimens. These must be cut into thin slices or sections in order to make them suitable for microscopical examination. Methods also exist for examination of surfaces: notably scanning electron microscopy (Echlin, 2009), atomic force microscopy (Vesenka et al., 1995), and in vivo light microscopic examination of mucosal surfaces for clinical diagnosis (Bornhop et al. 1999; Goetz et al. 2013). Preparative methods for such techniques are outside the scope of this book.

Freehand sections, cut with a razor, are rarely used in animal histology but are still sometimes employed for botanical material. Though some expertise is necessary, sectioning in this way has the advantage of requiring little in the way of time or special equipment. In a variant of the freehand technique, single sections of animal tissues are obtained by fixing a specimen with cyanoacrylate glue to either a glass slide or a cellulose acetate sheet, and then shaving it off with an inclined razor blade (Troyer et al., 2002; Dobkin and Troyer, 2003). When sections of human or animal tissues are needed in a hurry, frozen sections are commonly used. The cryostat, a microtome mounted in a freezing cabinet, may be used for cutting thin (5–10 µm) sections of either fixed or unfixed tissue. Each frozen section is collected from the knife onto a glass slide or coverslip, and does not thaw until it is removed from the cryostat cabinet. A traditional freezing microtome, with which the sections thaw as they are being cut, is used when thick (50–100 µm) sections are needed, especially in neuroscience research. Another advantage of cutting frozen sections, aside from speed, is the preservation of lipids, most of which are dissolved out during the course of dehydration and embedding in paraffin or plastic. A vibrating microtome (Vibratome) can cut thick (50–100 µm) sections of unfixed, unfrozen specimens. The blade of this instrument passes with a sawing motion through a block of tissue immersed in an isotonic saline solution. The cutting process is much slower than with other types of microtome, so it is not feasible to prepare large numbers of sections. Vibratome sections of fixed material are similar to sections cut with a freezing microtome, but they do not contain holes or other artifacts associated with ice crystal formation.

When the preservation of lipids or of heat-labile substances such as enzymes is not important, fixed specimens are dehydrated, cleared (which means, in this context, equilibrated with a solvent that is miscible with paraffin), infiltrated with molten paraffin wax, and finally, embedded (blocked out) as the wax cools and solidifies. Paraffin sections are most commonly cut on a rotary microtome, though a rocking microtome or a sledge microtome may also be used. The sections come off the knife in ribbons, and with sufficient skill it is possible to obtain serial sections as little as 4 µm thick through the whole block of tissue. The traditional embedding medium for large specimens was cellulose nitrate. Materials sold for the purpose were commonly called nitrocellulose, celloidin or low-viscosity nitrocellulose (LVN); these embedding media are seldom used today (see Chapter 4, Sections 4.1.3, 4.1.4). Various synthetic resins (plastics) are also used as embedding media for light microscopy, though their main application is in the cutting of extremely thin sections for examination in the electron microscope (EM). Resin-embedded tissue is usually sectioned with an ultramicrotome, using a glass or diamond knife. Semithin sections (0.5–1.0 µm), suitably stained for LM, are valuable for comparison with the much thinner sections used in EM studies. The LM provides greater resolution of detail in plastic- embedded sections than in paraffin sections, but the latter are more easily stained in contrasting colours. Larger resin-embedded objects are sectioned with a heavy-duty paraffin microtome and a tungsten carbide knife.

The optical contrast in a thin specimen is determined partly by its intrinsic properties but largely by the way in which it has been treated. If the specimen is not stained, contrast will be greatest when the mounting medium has a refractive index substantially different from that of the specimen. The visibility of a transparent specimen can be increased, at the expense of resolution, by defocusing the condenser of the microscope and by reducing the size of the substage diaphragm. Differences in refractile properties are emphasized in the phase contrast and the differential interference contrast (Nomarski) microscopes. These instruments are valuable for the study of living cells, such as those grown in tissue culture. With video-enhanced contrast otherwise inconspicuous features are enhanced by manipulation of electronically acquired images (see Shotton, 1993; Diaspro, 2002).

In histology, the natural refractility of a tissue is usually deliberately suppressed by the use of a mounting medium with a refractive index close to that of the anhydrous material constituting the section (approximately 1.53). Almost all the contrast is produced artificially by staining.

Fluorescence is the property exhibited by substances that absorb light of short wavelength such as ultraviolet or blue and emit light of longer wavelength, such as green, yellow, or red. The phenomenon can be observed with a fluorescence microscope in which arrangements are made for the emitted (longer wavelength) light to reach the eye while the exciting (shorter wavelength) light does not. Fluorescing materials therefore appear as bright objects on a dark background. The fluorescence microscope can be used to observe autofluorescence due to substances naturally present and secondary fluorescence produced by chemical modification of the specimen. The fluorescence of living cells arises from mitochondria and lysosomes (Andersson et al., 1998). Autofluorescence is due to various endogenous compounds, including flavoproteins, lipofuscin pigment, and elastin. Fluorescent compounds are also formed in tissues by chemical reactions between some fixatives and proteins (Collins and Goldsmith, 1981). See Section 1.4 for a brief introduction to fixation, and Chapter 2 for more information. Fixative-induced secondary fluorescence is often called autofluorescence. Any intrinsic fluorescence of a tissue is likely to interfere with the interpretation of secondary fluorescence. Various physical and chemical treatments can be used to suppress unwanted autofluorescence and fixative-induced fluorescence before or even after applying fluorescent reagents to sections, smears or cell cultures (see Kiernan, 2002a). Some of these are summarized in Table 1.1. These tricks for suppressing autofluorescence work in different ways, and some can interfere with techniques used to generate desirable secondary fluorescence. The references in the table should be consulted before applying the treatments to sections or cells used for diagnosis or research.

In confocal microscopy, the field is scanned, usually by a laser, to provide images of optical sections through thick specimens (see Diaspro, 2002; Hoppert, 2003). The images are derived from fluorescence (or, much less frequently, from reflected light). Optical sections are obtained by placing a pinhole in the light path between the objective and the detector. With a small enough pinhole, only the light emitted from an extremely thin layer within the specimen will reach the detector. With larger pinholes the thickness of the optical section increases. Images in different planes are recorded electronically, and synthesized to provide either three-dimensional pictures or flat pictures of selected objects that are too thick or tortuous to be seen in a single focal plane. Images from a confocal microscope are stored as files on a computer disk, and the contrast and other features can be manipulated prior to the production of a physical picture (see Wingate, 2002).

In multi-photon microscopy, the specimen is scanned by a pulsed high-power laser with a wavelength about double the absorption maximum of the fluorochrome that is to be excited. The pulses typically last 10-13s and are at 10-5s intervals. When two lower energy (longer wavelength) photons impinge on a fluorochrome molecule almost simultaneously (within 10-18s), their energies summate and stimulate the fluorescent emission of a photon. For example, a dye that absorbs blue light may be stimulated by a near-infrared laser to emit yellow light. This type of confocal microscopy allows the capture of optical sections thinner than those obtainable with single- photon confocal microscopy, and there is less photobleaching and less unwanted background fluorescence (Fuseler et al., 2011).

1.2. Staining and histochemistry

The histologist stains sections in order to see structural details. The histochemist, on the other hand, seeks to determine the locations of known substances within the structural framework. The disciplines of histology and histochemistry overlap to a large extent, but one consequence of the two approaches is that the staining techniques used primarily for morphological purposes are sometimes poorly understood in chemical terms. It is desirable to demonstrate structural components by 'staining' for substances they are known to contain, but many valuable empirically derived histological techniques are not based on well understood chemical principles. Nevertheless, the affinities of stains for particular components of cells and tissues depend on the chemical and physical properties, and often can be predicted from features easily seen in the structural formulae of dyes (Dapson and Horobin, 2013).

1.3. Some physical considerations

The intelligent handling of microscopical preparations requires familiarity with the physical properties of several materials that are used in almost all techniques. All too often, the beginner will ruin a beautifully stained section by forgetting that two solvents are immiscible, or by leaving the slides overnight in a liquid that dissolves the coloured product. The following remarks relate mainly to sections mounted on slides, but they apply also to blocks of tissue, smears, films, whole mounts, and free-floating sections.

Water is completely miscible with the common alcohols (methanol, ethanol, isopropanol). Water is immiscible with xylene, benzene, chloroform, and other non-polar solvents. These non-polar liquids, which are called clearing agents, are miscible with the alcohols in the absence of water. Melted paraffin wax and the resinous mounting media (Canada balsam, Cytoseal, DPX, Entellan, Permount, Xam, etc.) are miscible with the clearing agents but not with the alcohols or with water. One resinous mounting medium, euparal, is notable for being miscible with alcohol as well as with xylene. Because of these properties of the common solvents, a specimen must be passed through a series of liquids during the course of embedding, staining, and mounting for examination.

For example, a piece of tissue removed from an aqueous fixative, such as a formaldehyde solution, must pass through a dehydrating agent (such as alcohol) and a clearing agent (such as chloroform) before it can be infiltrated with paraffin wax. Ribbons of paraffin sections are floated on warm water, which removes wrinkles, and mounted on glass slides. A thin layer of a suitable adhesive (Chapter 4) may be interposed between the slide and the sections, but this is not always necessary. The slides must then be dried thoroughly in warm air before being placed in a clearing agent, usually xylene, to dissolve and remove the wax. The slides now bear sections of tissue that are equilibrated with the clearing agent. Passage through alcohol (or any other solvent miscible with both xylene and water) must precede immersion of the slides in water. Sudden changes are avoided if possible, so a series of graded mixtures of alcohol with water is used. Most staining solutions and histochemical reagents are aqueous solutions. If a permanent mount in a resinous medium is required, the slides carrying the stained sections must be dehydrated, without unintentionally removing the stain, in alcohol or a similar solvent, cleared (usually in xylene), and, finally, mounted by applying the resinous medium and a coverslip.

Several synthetic resins are used as embedding media. In most procedures the specimen is first infiltrated with a mixture of monomer and a catalyst at room temperature, and then moved to an oven (60 °C) to initiate polymerization. Most monomers are miscible with ethanol or other organic solvents. Some of the polymers are similarly soluble; others can only be removed from the sections by reagents that break covalent bonds in the matrix of resin.