Molecular Biology of the Cell - Volume 1 - Alberts B., Bray D., Lewis J., Raff M., Roberts K., Watson J. 1994

Introduction to Cell Biology
How Cells Are Studied
Microscopy

Cells are extremely small and complex: it is difficult to resolve their Structure, hard to determine their molecular composition, and even harder to establish how their individual components function. A variety of experimental Methods have been developed to study cells, and their capabilities define The Scope of our knowledge in this field. Advances in Cell biology, including the most remarkable achievements of recent years, have typically relied on the application of new methodological approaches. Therefore, to understand cell biology, one must have some grasp of the relevant experimental methods.

In this chapter, we will briefly review the modern methods used to study cells. We will begin with those that allow us to study The Cell as a whole, and then turn to the analysis of its constituent macromolecules. Microscopy is our starting point, as cell biology began with light microscopy, which remains a highly effective research tool alongside more modern imaging devices based on electron beams or Other forms of radiation. From passive observation, we will gradually move to active intervention: we will look at how different cell types can be isolated from tissue while retaining their ability to grow, and how cells can be disrupted to isolate their Organelles and constituent macromolecules in pure form. Finally, we will outline the principles of Recombinant DNA technology, which has made it possible to isolate, sequence, and manipulate genes, and thus study their Mechanisms of action within the cell.

A typical animal cell is 10–20 μm in diameter, which is about five times smaller than the smallest particle visible to the naked eye. Only with the advent of improved light microscopes in the early nineteenth century was it established that all animal and plant Tissues are composed of individual cells. This discovery, formulated as the Cell Theory by Schleiden and Schwann in 1838, marks the birth of cell biology.

In addition to being extremely small, animal cells are colorless and translucent; thus, the discovery of their major internal structures depended on The Development of a variety of organic Dyes in the late nineteenth century. It was these dyes that provided the contrast necessary to resolve subcellular structures. A similar situation arose in the early 1940s, when the invention of the powerful Electron microscope demanded new methods for preserving and staining cells. Only after these techniques were developed did the full complexity of Cell Structure begin to emerge. At its core, microscopy as a methodology still relies on specimen preparation and the inherent capabilities of the microscope itself.

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Fig. 4-1. The sizes of cells and their components, along with the useful ranges of light and electron microscopes, plotted on a logarithmic scale. In microscopy, the following units of length are commonly used: μm (micrometer) 10^-6 m, nm (nanometer) - 10^-9 m, Å (angstrom) - 10^-10 m.

Table 4-1. Major Milestones in the history of light microscopy

Fig. 4-2. Interference of light waves. If two light waves are in phase, the amplitude of the resulting wave increases, and thus the brightness increases. If two light waves are out of phase, they cancel each other out, producing a wave of reduced amplitude (and therefore reduced brightness).

Fig. 4-3. The interference effect that can be observed at high magnification at the edges of a solid object placed between a light source and an observer.

Figure 4-1 compares the resolution limits of modern light and electron microscopes.

The major milestones in the development of modern microscopy are listed in Table 4-1.

4.1.1. The light microscope can resolve objects separated by as little as 0.2 μm [2]

In general, radiation of a given wavelength can be used to study only those structures whose minimum dimensions are comparable to the wavelength of the radiation itself. This fundamental principle limits the capabilities of any microscope. The resolution limit of a light microscope is set by the wavelength of light, which for visible light ranges from 0.4 μm (violet) to 0.7 μm (deep red). Consequently, the smallest objects that can still be resolved under a light microscope are bacteria and mitochondria (which are about 0.5 μm wide). Smaller cellular components are distorted by effects caused by the wave nature of light. To understand The Nature of these effects, we must trace what happens to light waves as they pass through the lenses of a microscope.

Because of the wave nature of light, a beam of light does not travel in the perfectly straight path predicted by the laws of geometric optics. In reality, light waves travel through an optical system along many slightly different paths. Optical diffraction is caused by the interference of light waves whose paths through The Optical System differ slightly. If the light waves are precisely in phase—that is, the crest of one aligns with the crest of another, and the trough of one with the trough of another—they reinforce each other, and the brightness increases. On the other hand, if the waves are out of phase, they will cancel each other out (Fig. 4-2). Under high magnification, the shadow of a straight edge illuminated by light of a single wavelength will appear as a set of parallel lines, whereas a circular spot will appear as a set of concentric rings (Fig. 4-3). For the same reason, a single point appears in a microscope as a blurred disc, and two closely spaced point objects produce overlapping images that merge into one. Improving the precision of lens manufacturing cannot overcome this limitation, as it is imposed by the wave nature of light itself.

The limit of resolution—the minimum distance at which two objects can be distinguished as separate—depends on both the wavelength of light and the numerical aperture of the lens system used (Fig. 4-4). Under the most favorable conditions—using violet light (wavelength = 0.4 μm) and a numerical aperture of 1.4—the theoretical limit of resolution of the light microscope, about 0.2 μm, can be achieved. This limit was reached by microscope designers in the late nineteenth century (although it is rarely achieved in modern, mass-produced microscopes). And while an image can be magnified as much as desired, for example by projecting it onto a screen, a light microscope still cannot resolve two objects if they are separated by less than 0.2 μm: such objects will appear as a single entity.

The wave nature of light is not always an obstacle to studying cells; as we will see later, interference and diffraction can be exploited to study living, unstained cells. First, however, we must discuss the METHODS FOR PREPARING permanent cell specimens and how chemical stains can be used to enhance the visualization of cellular structures in such preparations.

Fig. 4-4. The path of light waves passing through a transparent specimen in a microscope, illustrating METABOLISM/2.html">THE CONCEPT OF numerical aperture and its relationship to the limit of resolution.

4.1.2. For microscopic examination, tissues are usually fixed and sectioned [2]

To prepare a permanent specimen that can be stained and observed under a microscope, cells must first be treated with a fixative to immobilize, kill, and preserve them. In chemical terms, fixation increases the accessibility of cells to dyes; cellular macromolecules are cross-linked, stabilizing and locking them in place. Some early fixation methods involved Treatment with acids or organic Solvents, such as alcohol. Modern methods typically use aldehydes, such as formaldehyde or glutaraldehyde, which form covalent bonds with the free amino groups of Proteins and thus cross-link neighboring molecules.

Most tissue specimens are too thick to allow individual cells to be examined directly at high resolution. Therefore, after fixation, tissues are usually cut into very thin slices (sections) using a microtome: an instrument with a very sharp metal blade that operates much like a bread slicer (Fig. 4-5). Sections from 1 to 10 μm thick are placed On the surface of a Glass slide. Because tissues are usually very soft and delicate even after fixation, they must be embedded in a supporting medium before sectioning. Typically, paraffin wax or a special resin is used as the embedding medium. In liquid form, these media infiltrate and surround the fixed tissue.

Figure 4-5. Preparing a section on a microtome after tissue embedding. The section is intended for examination under a light microscope. They harden upon cooling or through polymerization, forming a solid block that is easy to cut with a microtome.

There is a serious risk that fixation or embedding Procedures may damage The structure of cells or cellular macromolecules. This is why another sectioning method has been proposed to minimize this risk: rapid freezing. Here, fixation and embedding can be bypassed. Frozen tissue is simply cut using a cryostat—a specialized microtome housed in a cold chamber. Sections obtained this way avoid certain artifacts, yet they have their own disadvantages: while the individual structures of macromolecules, such as proteins, are well preserved during freezing, the overall STRUCTURE OF THE cell itself may be damaged. The next step after sectioning (by either method) is staining.

4.1.3. Different cell components can be stained in different ways [3]

Since the contents of most cells consist of about 70% Water, they contain almost no components capable of obstructing the passage of light rays. Consequently, in their natural state, most cells remain virtually invisible under a conventional light microscope, even after fixation and sectioning. One way to visualize them is by staining the cells with dyes.

In the early 19th century, driven by the demand for dyes in the textile industry, organic chemistry experienced a highly productive period. It turned out that some of these dyes were also capable of staining biological tissues. To the surprise of researchers, some of these dyes exhibited a specific affinity for particular cellular components, such as The Nucleus or mitochondria, staining their internal structures and making them visible for microscopic study. Today, a wide range of organic dyes is known. Many of them have colorful names, such as malachite green, Sudan black, and Coomassie blue; each dye is characterized by an affinity for specific subcellular components. For example, the dye hematoxylin has an affinity for negatively charged molecules and therefore reveals the distribution of DNA and acidic proteins in cells. The chemical basis for the Specificity of many dyes remains unknown.

As cell chemistry advanced, the most efficient and selective staining methods were developed, particularly those allowing the differentiation of specific proteins or other cellular macromolecules. This, however, raised the issue of sensitivity. Since most macromolecules are present in cells in relatively low copy numbers, one or two dye molecules bound to a macromolecule might go unnoticed. One way to solve this problem was to increase the number of dye molecules associated with individual cellular molecules. Many Enzymes could be localized within cells based on their catalytic activity: when supplied with an adequate substrate, each enzyme molecule generated numerous molecules of a visible reaction product. An alternative approach to the sensitivity problem is The Use of fluorescence. In this case, specific dyes can be detected against a dark Background by the light they, and only they, emit when appropriately excited. We will proceed to explain this phenomenon next.

4.1.4. Specific molecules can be localized in cells using Fluorescence Microscopy [4]

Fluorescent dyes absorb light of one wavelength and emit light of another, longer wavelength. If such a substance is illuminated with light of a wavelength that matches the absorption spectrum of the dye, and a filter is then used to transmit only the wavelength emitted by the dye, the fluorescent molecule can be detected by its glow against a dark background. High emission intensity is a characteristic feature of these molecules. The use of fluorescent dyes for cell staining requires a specialized fluorescence microscope. This microscope is similar to a conventional light microscope, but the light from a powerful source passes through two sets of filters—one to filter the light before it reaches the specimen, and another to filter the light coming from the specimen. The first filter is selected to pass only the wavelength that excites a particular fluorescent dye, while the second filter blocks this excitation light and allows only the wavelength emitted by the fluorescing dye to reach the eyepiece (Fig. 4-6). Fluorescence microscopy is frequently used to detect specific proteins or other molecules that become fluorescent after covalent binding to fluorescent dyes. For example, fluorescent dyes can be coupled to antibody molecules, instantly turning them into highly specific and convenient staining Reagents that selectively bind to specific macromolecules on The surface of living cells or inside fixed cells (see Section 4.5.3). Two dyes are commonly used for this purpose: fluorescein, which emits an intense yellow-green fluorescence when excited by blue light, and rhodamine, which produces a deep red fluorescence when excited by yellow-green light (Fig. 4-7). By using both fluorescein and rhodamine for staining, the distribution of different molecules can be studied; the Two Types of molecules can be visualized in the microscope simply by switching between two sets of filters, each specific to one of the dyes (Fig. 4-8).

Figure 4-6. The Optical System of a modern fluorescence microscope consists of two selective filters and a dichroic (beam-splitting) mirror. The set of filters used to detect fluorescein fluorescence is shown here. For this type of microscope, It is important to have objective lenses with a high numerical aperture, since for a given magnification, the brightness of the image is proportional to the fourth power of the aperture (see also Fig. 4-4).

Figure 4-7. In fluorescence microscopy, two dyes are commonly used: fluorescein and tetramethylrhodamine, whose structures are shown in this figure. Fluorescein emits yellow-green light upon activation by light of the appropriate wavelength. Rhodamine emits red light. The colored portion of the molecule indicates THE POSITION OF the chemically reactive group; a covalent bond between the dye and a protein (or other molecule) is typically formed at this position. Today, several variants of these dyes are commercially available with Different types of reactive groups, allowing the dye to be targeted to either the SH groups or the NH₂ groups of a protein.

Next, we turn to a Discussion of important new methods that allow the use of fluorescence microscopy to analyze Changes in the concentration and distribution of specific macromolecules in living cells (Section 4.1.9).

4.1.5. Phase-contrast and interference microscopes allow the Study of Living cells [2]

The potential for specimen loss or damage during preparation has always been a concern for microscopists. Only way to overcome this problem is to study living cells without fixation or freezing. Microscopes equipped with specialized optical systems are highly useful for this purpose.

As light passes through a living cell, the phase of the light wave changes According to the cell's refractive index: light passing through relatively dense or thick Regions of the cell, such as the nucleus, is retarded, and its phase is shifted relative to the phase of light passing through relatively thin areas of the Cytoplasm. Both phase-contrast and interference microscopes utilize the interference effects resulting from the recombination of two sets of waves to generate an image of cellular structures (Fig. 4-9). Both types of light microscopy are widely used for observing living cells.

Figure 4-8. Fluorescence micrograph of a surface region of early Drosophila embryos, where microtubules were labeled with fluorescein-conjugated antibodies (left) and Actin filaments with rhodamine-labeled antibodies (center). In addition, chromosomes were labeled with a third dye that fluoresces only when bound to DNA (right). At this stage, all nuclei of the embryo reside in a common cytoplasm and are not separated by cell walls; they are in metaphase of mitosis. All three micrographs were taken from the same region of a fixed embryo using three different filter sets in a fluorescence microscope (see also Fig. 4-6). (Courtesy of Tim Carr).

Figure 4-9. Two ways to increase contrast in light microscopy. A. Stained regions of the cell reduce the amplitude of transmitted light waves of certain wavelengths. As a result, a colored image visible upon direct observation can be obtained. B. The amplitude of light waves passing through an unstained living cell remains virtually unchanged; therefore, many details cannot be seen by direct observation. Here, however, a phase shift of the transmitted light occurs—a phenomenon utilized in phase-contrast and interference microscopes to produce a high-contrast image.

Figure 4-10. A fibroblast in tissue culture observed using four different types of light microscopy. A. The image is obtained by transmitting light directly through the cell (bright-field microscopy). The remaining images are obtained using the methods discussed in the text: B—Phase-contrast microscopy; C—interference microscopy; D—dark-field microscopy. Simply swapping the optical components of most modern microscopes allows all four types of images to be obtained.

Figure 4-11. An image of unstained microtubules observed under an interference microscope. A. The original, unprocessed image; B. The image obtained after electronic Processing, which significantly increases contrast and reduces noise. Although the diameter of microtubules is only 0.025 µm, due to diffraction they appear as much thicker filaments. (Courtesy of Bruce Schnapp.)

The simplest way to discern details of cellular structure is to observe the light scattered by various Components of the cell. In a dark-field microscope, light rays from the illuminator are directed from the side, so that only scattered rays enter the microscope lenses. Consequently, the cell appears as a bright object against a dark background. Images of the same cell obtained by four different light microscopy methods are shown in Fig. 4-10.

One of the major advantages of phase-contrast, interference, and dark-field microscopy is The ability to observe cell movements during mitosis and migration. Cell movements are usually very slow and difficult to follow in real time. In these cases, time-lapse (cinemicrographic) or video recording is used. Successive frames are separated by time intervals, but when the recording is played back at normal speed, the real-time events appear accelerated.

4.1.6. Images can be enhanced or analyzed using electronic methods [5]

In recent years, video cameras and associated Image processing technologies have significantly expanded the capabilities of light microscopy. Their application has not only overcome the limitations of optical systems but also resolved issues arising from human physiology. These limitations are that:

1) the eye cannot detect very dim light;

2) the eye is unable to perceive small differences in light intensity against a bright background.

The first of these problems was overcome by attaching ultra-high-sensitivity video cameras (similar to those used for night-time filming) to the microscope. This made it possible to observe cells over long periods at low light levels, avoiding prolonged exposure to bright light (or heat). Image intensification systems are particularly important for studying fluorescent molecules in living cells.

Because the image is generated by a video camera as electronic signals, it can be converted into digital signals, sent to a computer, and then further processed to extract hidden information. These and similar image processing techniques make it possible to compensate for optical defects in microscopes and practically reach the limit of resolution. Moreover, using modern video systems, contrast can be enhanced to such an extent that the eye's limitations in detecting small variations in light intensity are overcome. Although this process amplifies random background variations in the optical system, such 'noise' can be removed by Special Methods. Thus, thanks to modern approaches, we are now able to analyze transparent objects that were previously indistinguishable from the background.

The high contrast achievable with computer-assisted interference microscopy allows the observation of even very small objects, such as individual microtubules (Fig. 4-11), whose diameter is less than one-tenth the wavelength of light (0.025 µm). Individual microtubules can also be seen using fluorescence microscopy (see Fig. 4-56). However, in both cases, diffraction effects are inevitable and significantly distort the image. As a result, the apparent diameter of the microtubules is exaggerated (0.2 µm), making it impossible to distinguish a single microtubule from a bundle of several. To resolve this, an electron microscope is required, whose resolution limit is shifted far beyond the wavelength of visible light.

Fig. 4-12. Electron microscopy of a thin layer of gold reveals individual atoms as distinct bright spots. The distance between adjacent gold atoms is about 0.2 nm. (Courtesy of Graham Hills.)

4.1.7. The electron microscope allows the Analysis of the Fine Structure of the cell [6]

The relationship between the wavelength of light and the limit of resolution (see Fig. 4-4) holds true for any form of radiation, whether light rays or electrons. In the latter case, however, the resolution limit is significantly lower. The wavelength of an electron decreases as its velocity increases. In an electron microscope operating at 100,000 V, the electron wavelength is 0.004 nm, and theoretically, the resolution of such a microscope is 0.002 nm. However, correcting aberrations in electromagnetic lenses is a much more difficult task than for glass lenses, so in practice, the resolution of modern electron microscopes is at best 0.1 nm (1 Å) (Fig. 4-12). Furthermore, the difficulties of specimen preparation and radiation damage significantly reduce the practical resolution, which for biological specimens is about 2 nm (20 Å) (i.e., about 100 times higher than that of a light microscope).

Some of the milestones in the development of electron microscopy are listed in Table 4-2.

The general design of a transmission electron microscope (TEM) resembles that of a light microscope, although the electron microscope is much larger and essentially upside down (Fig. 4-13). The radiation source is a cathode filament that emits electrons from the top of a cylindrical Column about two meters high. Because electrons are scattered by collisions with air molecules, a vacuum must be maintained within the column. The electrons emitted by the cathode filament are accelerated by a nearby anode and pass through a tiny aperture, forming an electron beam that travels down the column. Ring-shaped magnets are positioned at intervals along the column to focus the electron beam, much like glass lenses focus a beam of light in a light microscope. The specimen is introduced through an airlock into the vacuum of the column, directly into the path of the electron beam. As they pass through the specimen, some electrons are scattered depending on the density of the material in that region, while the remaining electrons are focused to form an image (similar to image formation in a light microscope) on a photographic plate or a phosphorescent screen.

4.1.8. Biological specimens must undergo special preparation for observation under an electron microscope [7]

The application of the electron microscope to biology has revealed a wealth of amazing structures within cells. But before these discoveries could be made, scientists had to work hard to develop new methods for embedding, sectioning, and staining tissues.

Table 4-2. Major milestones in The history of electron microscopy

Fig. 4-13. Schematic diagram of the Main Components of a light microscope, a transmission electron microscope, and a scanning electron microscope, highlighting the similarities in their design. In both types of electron microscopes, the specimen must be placed in a vacuum.

Fig. 4-14. Glutaraldehyde and osmium tetroxide are the most common fixatives used in electron microscopy. The two reactive aldehyde groups of glutaraldehyde allow it to cross-link various types of molecules via covalent bonds that displace hydrogen atoms (shown in color). Osmium tetroxide is reduced by many Organic compounds, forming cross-linked complexes with them. This property is especially valuable for cell membranes, as osmium tetroxide reacts with the C-C double bonds characteristic of many Fatty acids.

Fig. 4-15. Schematic diagram of a copper grid used to support thin specimen sections in a transmission electron microscope.

In an electron microscope, all specimens are exposed to a high vacuum, making it impossible to observe them in a living, wet state. Tissues are typically preserved by fixation—first with glutaraldehyde, which covalently cross-links protein molecules, and then with osmium tetroxide, which binds and stabilizes The Lipid Bilayer and proteins (Fig. 4-14). Because electrons have low penetrating power, specimens must be cut into extremely thin sections, from 50 to 100 nm thick (about 1/200 of the thickness of a single cell). Only such sections can be observed in an electron microscope. This also requires prior dehydration of the specimen and infiltration with monomeric resins that, upon polymerization, form a solid plastic block embedding the specimen; the block is then cut with a fine glass or diamond knife on a specialized microtome. The resulting sections contain no water or other volatile organic substances; these sections are placed on a small circular metal grid for observation under the microscope (Fig. 4-15).

The contrast in an electron microscope is determined by the atomic number of the elements in the specimen. The higher the atomic number, the more electrons are scattered, and the greater the contrast. Biological molecules are composed of atoms with very low atomic numbers (mainly oxygen, hydrogen, carbon, and nitrogen). To enhance contrast, specimens are stained with heavy metal salts, such as osmium, uranium, and lead, before and after sectioning. Cellular components are revealed with varying degrees of contrast depending on how heavily they are stained by these salts. Typically, Lipids are stained dark by osmium, allowing membranes to be visualized (Fig. 4-16).

In some cases, electron microscopy techniques can be used to localize specific macromolecules in thin sections. Certain cellular enzymes are detected after incubating specimens with substrates whose enzymatic reaction leads to the local deposition of an electron-dense precipitate (Fig. 4-17). In addition, labeled antibodies can be used to localize specific macromolecules if they are coupled to an indicator enzyme such as peroxidase or to an electron-dense marker, such as colloidal gold particles (see Section 4.5.3).

Fig. 4-16. Thin section of a grass ROOT tip. The Cell wall, nucleus, vacuoles, mitochondria, Endoplasmic reticulum, Golgi apparatus, and Ribosomes are easily distinguishable. (Courtesy of Brian Gunning.)

Fig. 4-17. Electron micrograph of a cell illustrating the localization of a specific enzyme (nucleoside diphosphatase) in the Golgi apparatus. A thin section of the cell was incubated with a substrate that forms an electron-dense precipitate under the action of the enzyme. (Courtesy of Daniel Friend.)

Fig. 4-18. Schematic diagram illustrating how easily one can draw erroneous Conclusions from studying individual thin sections. In this case, for example, we look at sections of a cell containing only a single branched mitochondrion. However, it appears as though most of the sections contain two or three separate mitochondria. Furthermore, in sections 4 and 7, the visualized mitochondrion might seem to be in The process of dividing. Serial sections allow the reconstruction of the actual three-dimensional shape.

4.1.9. The scanning electron microscope is used to obtain three-dimensional images of the surface [8]

Thin sections are essentially two-dimensional slices of tissue and do not allow one to determine the three-dimensional structure of cellular components. A three-dimensional image can be obtained by reconstructing hundreds of serial sections (Fig. 4-18), but this is a long and tedious process. Today, more direct methods for obtaining three-dimensional images have been developed. One of these is to examine the specimen under a scanning electron microscope (SEM), which is typically smaller and simpler than a transmission electron microscope. While a transmission electron microscope forms an image using electrons that pass through the specimen, a scanning electron microscope uses electrons that are scattered or emitted from the specimen's surface. In this case, the specimen must be fixed, dried, and coated with a thin film of heavy metal. The specimen is then scanned by a very narrow beam of electrons. The number of electrons scattered as successive points on the metallic surface are irradiated is measured. This value is used to control the intensity of a second beam that moves synchronously with the first, forming an image on a television screen. In this way, a single, cohesive, and highly magnified image is constructed.

Fig. 4-19. Scanning electron micrograph of stereocilia arranged like organ pipes on the surface of Hair cells in the Inner ear. (Courtesy of R. Jackobs, A.J. Hudspeth.)

Fig. 4-20. Electron micrograph of individual Myosin protein molecules (platinum-shadowed). Myosin is the major component of the contractile apparatus of muscle; here it can be seen to consist of two globular regions joined to a common rodlike tail. (Courtesy of Arthur Elliot.)

Fig. 4-21. Schematic representation of the method for preparing a replica of a specimen surface (metal shadowing). Note that the thickness of the metal layer is determined by the surface contour of the original specimen.

Scanning electron microscopy provides a great depth of field; furthermore, because the extent of electron scattering is determined by the angle of the surface relative to the beam, the image displays alternating light and dark areas that create a three-dimensional effect (Fig. 4-19). However, this method is only applicable for surface studies, and its resolution is relatively low (about 10 nm, with an effective magnification of approximately 20,000 times). Consequently, this technique is virtually useless for studying subcellular organelles and is used almost exclusively for examining whole Cells and Tissues.

Using conventional thin sections, tilting them, and photographing them at different angles, a simulated three-dimensional image can also be obtained in a standard transmission electron microscope. When viewing the resulting stereo pair of images through stereo glasses, a pseudo-three-dimensional image is perceived. The thickness of specimens studied by this method is limited by the penetrating power of the electrons, which depends on their energy. Advancements in this technique led to the development of high-voltage electron microscopes with accelerating voltages of up to 1 million volts (compared to 100,000 volts in conventional TEMs). These giant instruments allow sections several micrometers thick to be studied by transmission electron microscopy.

4.1.10. Metal shadowing is used to study surface details in the transmission electron microscope [9]

The transmission electron microscope can be used to study the surface of a specimen at very high magnification, allowing individual macromolecules to be observed. As in scanning electron microscopy, a thin film of heavy metal, such as platinum, is evaporated onto the dried specimen. The metal is sprayed from a specific angle so that the deposited film is thicker in some places than in others. This process, known as metal shadowing, creates a shadow effect that gives the image a three-dimensional appearance.

Specimens prepared in this way can be small and thin enough for the electron beam to penetrate them; for example, individual molecules, Viruses, and cell walls can be analyzed in this manner (Fig. 4-20). For thicker specimens, however, the organic material of the cell must be dissolved away after shadowing, leaving only a thin metal cast, or replica, of the surface. This replica is then reinforced with a carbon film, after which it can be mounted on a grid and examined in a conventional electron microscope (Fig. 4-21).

4.1.11. Freeze-fracture and freeze-etching electron microscopy provide a unique opportunity to observe the internal structure of the cell [10]

In cell biology, two Methods based on preparing mechanical replicas have been particularly successful. One of these, the freeze-fracture electron Microscopy Technique, provides a way to examine the internal Structure of Cell membranes. Cells are frozen at liquid nitrogen Temperature (-196°C) in the presence of a cryoprotectant (antifreeze) to prevent damage from ice crystal formation. The frozen block is then fractured with a cold knife blade. The fracture plane often splits the hydrophobic interior of the lipid bilayer, exposing the internal faces of cell membranes. The resulting fracture face is shadowed with platinum, the organic material is dissolved away, and the replicas are examined in an electron microscope (Fig. 4-21). Such replicas are studded with small bumps—intramembrane particles—which represent large Membrane Proteins. This method is extremely useful and effective for analyzing the distribution of these proteins within the membrane plane (Fig. 4-22).

Fig. 4-22. Electron micrograph of a plant cell chloroplast thylakoid membrane prepared by freeze-fracture. The photosynthetic thylakoid membranes are stacked in multiple layers. The fracture plane jumps from one layer to another, passing through the middle of each lipid bilayer. Intramembrane proteins, which are abundant within the bilayer, are exposed and, after shadowing, appear as intramembrane particles in this platinum replica. The largest particle visible in the membrane is a multi-protein complex forming Photosystem II. (Courtesy of L. F. Staehelin.)

Fig. 4-23. Freeze-etch electron microscopy. The frozen specimen is fractured with a knife (A). Then, by sublimating water under vacuum, the ice layer is reduced, thereby exposing the cell surface (B). After this, a replica of the still-frozen surface is prepared (as described in the legend to Fig. 4-21) and examined using a transmission electron microscope.

Fig. 4-24. Regularly arranged protein filaments in insect muscle. To obtain this image, muscle cells were frozen in liquid helium, fractured through the cytoplasm, and deeply etched. Then, a metal replica was prepared and examined at high magnification. (Courtesy of Roger Cooke, John Heuser).

Another important electron microscopy technique—freeze-etching—is used to study the outer surface of cells and membranes. In this method, cells are frozen at a very low temperature, and the frozen block is fractured with a knife blade. The ice surrounding the cells (and to a lesser extent inside them) is sublimated in a vacuum as the temperature is raised (a process known as freeze-drying) (Fig. 4-23). The etched regions of the cell are then shadowed (as shown previously) to prepare a platinum replica.

The freeze-etching method does not allow the use of cryoprotectants, because they are non-volatile and remain in the specimen as the water sublimates. To achieve high image quality, The formation of large ice crystals must be prevented. This is possible by rapid freezing of the specimen (at a freezing rate higher than 20°C per millisecond). One method of such rapid freezing involves using a special device that quickly brings the specimen into contact with a copper block cooled to –269°C by liquid helium. Particularly impressive results are obtained after deep etching of rapidly frozen cells. This technique reveals the internal structures of cells, demonstrating their three-dimensional Organization with exceptional clarity (Fig. 4-24).

Since in this case it is the metal-shadowed replicas rather than the specimens themselves that are observed in the microscope under vacuum, freeze-fracture and freeze-etch methods can be used to study frozen, unfixed cells, thereby eliminating the risk of artifacts caused by fixation.

4.1.12. Negative Staining and Cryoelectron Microscopy Provide High Resolution for Macromolecular Analysis

By using heavy metal shadowing for contrast, isolated macromolecules such as DNA or large Proteins can be observed in the electron microscope (see Fig. 4-20), and after negative staining, even the smallest details can be resolved. In preparing specimens for negative staining, the molecules under study are applied to a thin carbon film (which is virtually transparent to electrons), which is then wetted with a concentrated solution of heavy metal salts, such as uranyl acetate. After the specimen dries, a thin film of heavy metal salts uniformly covers the carbon support, except for the areas occupied by the adsorbed macromolecules. The macromolecular material is more permeable to electrons than the surrounding areas covered with heavy metal salts, resulting in a reversed or negative image of the molecule. Negative staining is particularly effective for observing large macromolecular aggregates (viruses, ribosomes) or for studying the subunit structure of protein filaments (Fig. 4-25).

Fig. 4-25. Electron micrographs of negatively stained actin filaments. The diameter of each of these filaments is approximately 8 nm. Close examination reveals that actin filaments consist of two helically wound chains of globular actin molecules. (Courtesy of Roger Craig.)

Negative staining and shadowing techniques provide high-contrast surface images of small macromolecular aggregates, but the resolution of these methods is limited by the size of the metal particles forming the shadow or the dye molecules consisting of heavy metal salts, which only roughly outline the surface of the molecule or macromolecular assembly. Today, however, even the internal details of three-dimensional structures, such as viruses, can be observed at high resolution. This is achieved using cryoelectron microscopy, where a very thin (approximately 100 nm) rapidly frozen layer of a wet specimen is placed on a microscope grid. Using a special holder, the hydrated specimen is maintained at –160°C in the microscope vacuum. In this way, the material can be observed almost directly: without fixation, staining, or drying. The homogeneity of the vitrified aqueous layer and the use of underfocused phase contrast make it possible to obtain remarkably clear images of such unstained specimens (Fig. 4-26).

Regardless of the methods used, individual protein molecules yield faint and poorly defined images in the electron microscope. Attempts to extract information by extending the exposure time or increasing the intensity of the illuminating beam are futile, as this leads to the destruction of the specimen. To analyze details of molecular structure, information from many molecules must be combined to avoid random errors inherent in individual images. This approach is suitable for studying viruses or protein filaments, whose individual subunits are arranged as regularly repeating elements; it is also applicable to any substances that can be arranged in a two-dimensional crystalline lattice, where A large number of molecules maintain the same orientation or are separated by identical intervals. Using electron micrographs of such oriented structures, image processing techniques can be applied to calculate an average image of individual molecules, revealing details that were obscured by the random 'noise' of the original photograph.

Fig. 4-26. Semliki Forest virus in a thin layer of unstained vitrified water (cryoelectron microscopy at –160°C). Using electronic image processing of micrographs, a high-resolution three-dimensional image can be obtained. (Courtesy of Jacques Dubochet; see also S.D. Fuller, Cell, 48, 923-934, 1987.)

Image reconstructions using this method allow the detailed structure of the virus envelope to be resolved to 3.5 nm, and the details of individual macromolecular shapes can be studied at a resolution of 0.5 nm (5 Å). Yet even the most sophisticated electron microscopy techniques cannot fully describe molecular structure, because atoms in molecules are separated by distances of about 0.1–0.2 nm. To study the molecular structure of macromolecules at the atomic level, other methods are required—methods that use X-rays instead of electrons.

4.1.13. The Detailed Structure of Molecules Forming a Crystalline Lattice Can Be Calculated from Diffraction Patterns [12]

X-rays, like light, are a form of electromagnetic radiation, but because their wavelength is much shorter, they can resolve much smaller details. However, unlike visible light or a beam of electrons, X-rays cannot be focused to produce a conventional image after passing through a specimen. Nevertheless, the structure of a specimen can be revealed using X-Ray Diffraction.

First, let us consider a single object (for example, an individual molecule) placed in the path of any radiation whose wavelength is shorter than the size of the object. The object will scatter some of the radiation. The scattered radiation can be viewed as a set of overlapping waves, each reflected from different PARTS OF THE object. When these waves overlap, they interfere with one another, producing a distribution of radiation known as a diffraction pattern. The diffraction pattern can be recorded on a photographic plate placed at some distance from the object, or represented by The amount of scattered radiation reflected by the object in different directions (Fig. 4-27). The shape of the diffraction pattern is determined by the structure of the object. Conversely, from a complete Description of the diffraction pattern, one can theoretically calculate the structure of the object. Experience shows that the diffraction pattern from a single molecule is far too weak and noisy to be used as a starting point for theoretical analysis.

Suppose that many identical objects are arranged in a crystalline lattice and a beam of radiation is directed at them (Fig. 4-28). In this case, the total amount of scattered radiation is much higher. However, the radiation scattered by one molecule will interfere with the radiation scattered by other molecules. In certain directions, the individual scattered rays will reinforce each other, forming a bright spot on the diffraction pattern.

Fig. 4-27. Scattering of radiation by a single object whose dimensions are comparable to the wavelength of the radiation. Radiation incident on the object is scattered in different directions and with different intensities. The intensity of the radiation scattered in a given direction depends on the interference of the radiation scattered by different parts of the object. The resulting scattering intensity in all possible directions can be judged by the number of colored rays in the diagram.

Fig. 4-29. Part of an X-ray diffraction pattern of a protein crystal. This particular crystal was used to determine the positions of atoms in the molecule of the proteolytic enzyme Trypsin. (Courtesy of Robert Stroud.)

Figure 4-28. Scattering of radiation by a crystal. If many identical objects are arranged in a crystal lattice, the radiation scattered by each object interferes with the radiation scattered by other objects. Individual reflected rays are scattered only in specific directions (depending on the spatial arrangement of the object in the lattice), forming bright spots. The intensity of a given bright spot depends on the intensity with which each individual object in the lattice would scatter radiation in that direction if examined in isolation, as shown in Fig. 4-27.

Figure 4-30. Crystals of the enzyme Glycogen phosphorylase as seen under a light microscope. (Courtesy of Robert Fletterick.) The diffraction pattern of a crystal lattice consists of many bright spots of varying intensity (Fig. 4-29). The relative intensity of the different spots in the diffraction pattern depends on the ability of the various objects in the lattice to scatter radiation. In fact, the intensity of a given spot is proportional to the intensity of the radiation that would be reflected in that direction from a single characteristic object. Thus, the position of the spots in the diffraction pattern depends on the arrangement of the object in the system, while their intensity provides information about the internal structure of a typical object. Moreover, such information is highly accurate and sufficient because it is obtained by combining the contributions of many equivalent sources. Indeed, using a fairly complete description of the diffraction pattern of such a lattice, one can often calculate the structure of the individual objects that make up the crystal lattice.

4.1.14. X-ray diffraction makes it possible to reveal the three-dimensional organization of atoms in molecules [13]

If diffraction patterns are to be used for molecular structure analysis, the diffracted radiation must have a wavelength shorter than the distance between atoms in the molecule. The wavelength of X-rays is about 0.1 nm (which corresponds to the diameter of a hydrogen atom), making this type of radiation ideal for analyzing the arrangement of individual atoms in molecules. This task cannot be accomplished even with the most advanced electron microscopes. A significant advantage of X-rays is their high penetrating power (higher than that of electrons). This makes thicker specimens suitable for analysis. Finally, since a vacuum is not required in this case, thick, water-containing specimens can be studied. Consequently, artifacts arising during specimen preparation are eliminated.

To achieve high resolution, highly ordered crystals are required (Fig. 4-30). As X-rays pass through the specimen, they are scattered by the electrons of the atoms that make up the sample. Therefore, large atoms with many electrons scatter X-rays more effectively than small atoms, so that C, N, O, and P atoms are detected much more reliably than H atoms; very heavy atoms are also known to scatter X-rays highly efficiently. The process of converting an X-ray diffraction pattern into a three-dimensional structure of atoms arranged in a molecule is highly complex. Deciphering diffraction patterns formed by large and disordered protein molecules was impossible before 1960 (Table 4-3). This Procedure requires locating and estimating the intensity of hundreds of thousands of spots, as well as the wave phases of each spot. In recent years, X-ray crystallography has become increasingly automated. Scattered X-rays are measured by sophisticated electronic detectors, which significantly speeds up the data collection process, while powerful computers perform the many necessary calculations. Currently, the most time-consuming step in such a study is obtaining suitable crystals of the macromolecules under investigation; it often takes years to find the optimal crystallization conditions. Despite these difficulties, X-ray crystallography has found wide application, as it remains the only method to date for determining the detailed arrangement of atoms in most molecules. With good crystals, the Protein Structure can be calculated at a resolution of 0.3 nm, revealing not only the Main Features of the polypeptide chain arrangement but also some finer details. With significant effort, high-quality crystals can be obtained, which in turn allows for a resolution of 0.15 nm and the Determination of the positions of almost all non-hydrogen atoms in the protein molecule. It is in this way that the structures of more than a hundred proteins and several small RNA and DNA molecules have been established to date.

Table 4-3. Major milestones in the development of X-ray crystallography and its application in the study of biological molecules

Conclusion

A variety of light microscopy techniques are available for observing cells. Stained and fixed cells can be viewed using conventional optics. The use of a fluorescence microscope and labeled antibodies allows for the localization of specific molecules within cells. Cells in their natural, living state are analyzed using phase-contrast, interference, or dark-field optics. Light microscopic studies of living cells are enhanced by electronic image processing, which significantly increases sensitivity and improves resolution.

Determining the detailed structure of membranes and organelles in cells is possible only at the high resolution provided by a transmission electron microscope. The transmission electron microscope is also used to determine the shape of individual macromolecules shadowed with heavy metals or negative staining. However, the precise Location of each atom in a molecule can only be determined after the molecules form large crystals. In this case, the complete three-dimensional structure of the molecule can be calculated using X-ray crystallography.