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Lodish H, Berk A, Zipursky SL, et al. Molecular Cell Biology. 4th edition. New York: W. H. Freeman; 2000.

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Molecular Cell Biology. 4th edition.

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Section 5.1Microscopy and Cell Architecture

The modern, detailed understanding of cell architecture is based on several types of microscopy. Because there is no one “correct” view of a cell, it is essential to understand the characteristics of the key cell-viewing techniques, the types of images they produce, and their limitations.

Schleiden and Schwann, using a primitive light microscope, first described individual cells as the fundamental unit of life, and light microscopy has continued to play a major role in biological research. The development of electron microscopes greatly extended the ability to resolve subcellular particles and has yielded much new information on the organization of plant and animal tissues. The nature of the images depends on the type of light or electron microscope employed and on the way in which the cell or tissue has been prepared. Each technique is designed to emphasize particular structural features of the cell. Figure 5-1 shows how a typical cell, the epithelial cell lining the small intestine, appears when viewed by three different microscopic techniques.

Figure 5-1. Views of the epithelial cells lining the small intestine, produced by three different microscopic techniques.

Figure 5-1

Views of the epithelial cells lining the small intestine, produced by three different microscopic techniques. (a) Scanning electron micrograph of the intestinal wall. The lumen, or cavity, of the intestine, is lined by a sheet of epithelial cells that rests (more...)

In this section, we focus on the most common application of light and electron microscopy — to visualize fixed, killed cells. Although this approach reveals much information, a critical question about such results is how true to life is the image of a biological specimen that has been fixed, stained, and dehydrated before examination? Thus we also consider some of the refinements that allow microscopy of unaltered or less altered specimens.

Light Microscopy Can Distinguish Objects Separated by 0.2 μm or More

The compound microscope, the most common microscope in use today, contains several lenses that magnify the image of a specimen under study (Figure 5-2a). The total magnification is a product of the magnification of the individual lenses: if the objective lens magnifies 100-fold (a 100X lens, the maximum usually employed) and the eyepiece magnifies 10-fold, the final magnification recorded by the human eye or on film will be 1000-fold.

Figure 5-2. The optical pathway in a modern compound optical microscope.

Figure 5-2

The optical pathway in a modern compound optical microscope. (a) The specimen is usually mounted on a transparent glass slide and positioned on the movable specimen stage of the microscope. Light from a bright source is focused by the condenser lenses (more...)

However, the most important property of any microscope is not its magnification but its resolving power, or resolution — its ability to distinguish between two very closely positioned objects. Merely enlarging the image of a specimen accomplishes nothing if the image is blurry. The resolution of a microscope lens is numerically equivalent to D, the minimum distance between two distinguishable objects; the smaller the value of D, the better the resolution. D depends on three parameters, all of which must be considered in order to achieve the best possible resolution: the angular aperture, α, or half-angle of the cone of light entering the objective lens from the specimen; the refractive index, N, of the air or fluid medium between the specimen and the objective lens; and the wavelength, λ, of incident light: D = (0.61λ) ÷ (N × sin α). Decreasing the value of λ or increasing either N or α will decrease the value of D and thus improve the resolution. Note that the magnification is not part of this equation.

The angular aperture, α, depends on the width of the objective lens and its distance from the specimen (Figure 5-2b). Moving the objective lens closer to the specimen increases the angle α and thus sin α, and therefore reduces D (i.e., increases the resolution). Intuitively, one can recognize that increasing α allows a greater fraction of the light emanating from the specimen to enter the objective lens. The refractive index N is a measure of the degree to which a medium bends a light ray that passes through it; the refractive index of air is defined as 1.0. Use of immersion oil, which has a refractive index of 1.5, is a simple way to reduce D by 33 percent. An intuitive explanation for this improvement is that a medium with a higher refractive index than air, if placed between the specimen and the objective lens, will “bend” more of the light emanating from the specimen such that it goes into the lens. Finally, the shorter the wavelength of incident light, the lower will be the value of D and the better the resolution.

Due to limitations on the values of α, λ, and N, the limit of resolution of a light microscope using visible light is about 0.2 μm (200 nm). No matter how many times the image is magnified, the microscope can never resolve objects that are less than ≈0.2 μm apart or reveal details smaller than ≈0.2 μm in size. This is true because the maximum angular aperture for the best objective lenses is 70° (sin 70° = 0.94). With the visible light of shortest wavelength (blue, λ = 450 nm) and with an immersion oil (N  =  1.5) above the sample, then

Image ch5e1.jpg

or about 0.2 μm.

Despite this limit of resolution, the light microscope can be used to track the location of a small bead of known size to a precision of only a few nanometers! If we know the precise size and shape of an object — say, a 5-nm sphere of gold — and if we use a video camera to record the microscopic image as a digital image, then a computer can calculate the position of the center of the object to within a few nanometers. This technique has been used, to nanometer resolution, for tracking the movement of gold particles attached via antibodies to specific proteins on the surface of living cells.

Samples for Light Microscopy Usually Are Fixed, Sectioned, and Stained

Specimens for light microscopy are commonly fixed with a solution containing alcohol or formaldehyde, compounds that denature most proteins and nucleic acids. Formaldehyde also cross-links amino groups on adjacent molecules; these covalent bonds stabilize protein-protein and protein – nucleic acid interactions and render the molecules insoluble and stable for subsequent procedures. Usually the sample is then embedded in paraffin or plastic and cut into thin sections of one or a few micrometers thick (Figure 5-3). Alternatively, the sample can be frozen without prior fixation and then sectioned; this avoids the denaturation of enzymes by fixatives such as formaldehyde.

Figure 5-3. Preparation of tissues for light microscopy.

Figure 5-3

Preparation of tissues for light microscopy. A piece of fixed tissue is dehydrated by soaking it in alcohol-water solutions, then in pure alcohol, and finally in a solvent such as xylene. The specimen is next placed in warm liquid paraffin, which is allowed (more...)

Since the resolution of the light microscope is ≈0.2 μm and mitochondria and chloroplasts are ≈1 μm long (about the size of bacteria), theoretically one should be able to see these organelles. However, most cellular constituents are not colored and absorb about the same degree of visible light, so that they are hard to distinguish under a light microscope unless the specimen is stained. Thus the final step in preparing a specimen for light microscopy is to stain it, in order to visualize the main structural features of the cell or tissue. Many chemical stains bind to molecules that have specific features. For example, hematoxylin binds to basic amino acids (lysine and arginine) on many different kinds of proteins, whereas eosin binds to acidic molecules (such as DNA, and aspartate and glutamate side chains). Because of their different binding properties, these dyes stain various cell types sufficiently differently that they are distinguishable visually. Two other common dyes are benzidine, which binds to heme-containing proteins and nucleic acids, and fuchsin, which binds to DNA and is used in Fuelgen staining.

If an enzyme catalyzes a reaction that produces a colored or otherwise visible precipitate from a colorless precursor, the enzyme may be detected in cell sections by their colored reaction products. This technique is called cytochemical staining (Figure 5-4).

Figure 5-4. Cytochemical staining.

Figure 5-4

Cytochemical staining. Light micrograph of a cross section of human skeletal muscle stained for succinate dehydrogenase, an enzyme found only in mitochondria. At this low magnification the stained mitochondria appear as purple dots; in skeletal muscle (more...)

Fluorescence Microscopy Can Localize and Quantify Specific Molecules in Cells

Perhaps the most versatile and powerful technique for localizing proteins within a cell by light microscopy is fluorescent staining of cells and observation in the fluorescence microscope. A chemical is said to be fluorescent if it absorbs light at one wavelength (the excitation wavelength) and emits light (fluoresces) at a specific and longer wavelength. Most fluorescent dyes emit visible light, but some (such as Cy5 and Cy7) emit infrared light. In modern fluorescence microscopes, only fluorescent light emitted by the sample is used to form an image; light of the exciting wavelength induces the fluorescence but is then not allowed to pass the filters placed between the objective lens and the eye or camera (Figure 5-5).

Figure 5-5. The optical pathway in an epi-fluorescence microscope.

Figure 5-5

The optical pathway in an epi-fluorescence microscope. Light from a multiwavelength source moves through an excitation filter, which allows only the desired wavelength of exciting radiation to pass. This radiation is reflected downward by the dichroic (more...)

Revealing Specific Proteins in Fixed Cells

Four very useful dyes for fluorescent staining are rhodamine and Texas red, which emit red light; Cy3, which emits orange light; and fluorescein, which emits green light. These dyes have a low, nonspecific affinity for biological molecules, but they can be chemically coupled to purified antibodies specific for almost any desired macromolecule. When a fluorescent dye – antibody complex is added to a permeabilized cell or tissue section, the complex will bind to the corresponding antigens, which then light up when illuminated by the exciting wavelength, a technique called immunofluorescence microscopy (Figure 5-6). By staining a specimen with two or three dyes that fluoresce at different wavelengths, multiple proteins can be localized within a cell, as illustrated in the chapter opening figure and in Figure 5-1c.

Figure 5-6. Fluorescence micrograph showing the distribution of long actin fibers in a cultured fibroblast cell.

Figure 5-6

Fluorescence micrograph showing the distribution of long actin fibers in a cultured fibroblast cell. A fixed human skin fibroblast was permeabilized with a detergent and stained with a fluorescent anti-actin antibody before viewing. [Courtesy of E. Lazarides.] (more...)

Revealing Specific Proteins in Living Cells

Fluorescence microscopy can also be applied to live cells. For example, purified actin may be chemically linked to a fluorescent dye. Careful biochemical studies have established that this “tagged” molecule is indistinguishable in function from its normal counterpart. If the tagged protein is microinjected into a cultured cell, the endogenous cellular and injected tagged actin monomers copolymerize into normal long actin fibers. This technique can also be used to study individual microtubules within a cell.

Another technique for detecting specific proteins within living cells takes advantage of green fluorescent protein (GFP), a naturally fluorescent protein found in the jellyfish Aequorea victoria. The bioluminescence of this organism, which radiates a green fluorescence, is due to GFP. This 238-aa protein contains serine, tyrosine, and glycine residues whose side chains have spontaneously reacted with one another to form a fluorescent chromophore. By recombinant DNA techniques discussed in Chapter 7, the GFP gene can be introduced into living cultured cells or into specific cells of an entire animal. Because the introduced gene will express GFP, the cells will emit a green fluorescence when irradiated; this GFP fluorescence can be used to localize the cells within a tissue.

Image biotech.jpgAlternatively, the gene for GFP can be fused to the gene for another protein of interest, producing a recombinant DNA encoding one long chimeric protein that contains the entirety of both proteins. Cells in which this recombinant DNA has been introduced will synthesize this chimeric protein, whose green fluorescence will reveal the subcellular localization of the protein. Figure 5-7 illustrates how this technique can demonstrate changes in the localization of a protein within a living cell following treatment with a particular hormone.

Figure 5-7. Use of green fluorescent protein (GFP) to localize GLUT4, a glucose transport protein, within living fat cells.

Figure 5-7

Use of green fluorescent protein (GFP) to localize GLUT4, a glucose transport protein, within living fat cells. Cells were engineered to express a chimeric protein whose N-terminal end corresponded to the GLUT4 sequence, followed by the entirety of the (more...)

Determining the Intracellular Concentration of Ca2+ and H+ Ions

Changes in the cytosolic concentration of Ca2+ ions or pH frequently signal changes in cellular metabolism. The Ca2+ concentration in the cytosol of resting cells, for instance, is about 10−7 M. Many hormones or other stimuli cause a rise in cytosolic Ca2+ to 10−6 M; this, in turn, causes changes in cellular metabolism, such as contraction of muscle (Chapter 18).

The fluorescent properties of certain dyes, such as fura-2, facilitate measurement of the concentration of free Ca2+ in the cytosol. This dye contains five carboxylate groups that form ester linkages with ethanol. The resulting fura-2 ester is lipophilic and can diffuse from the medium across the plasma membrane into cells. Within the cytosol, esterases hydrolyze fura-2 ester yielding fura-2, whose free carboxylate groups render the molecule nonlipophilic, so it cannot cross cellular membranes and remains in the cytosol. Each fura-2 molecule can bind a single Ca2+ ion but no other cellular cation, and the amount of fura-2 bound to Ca2+ is proportional, over a certain range, to the Ca2+ concentration. The fluorescence of fura-2 at one particular wavelength is enhanced when Ca2+ is bound, and the fluorescence is proportional to the Ca2+ concentration. At another wavelength the fluorescence of fura-2 is the same whether or not Ca2+ is bound and provides a measure of the total amount of fura-2 in the segment of the cell. By examining cells continuously in the fluorescence microscope and measuring rapid changes in the ratio of fura-2 fluorescence at these two wavelengths, one can quantify rapid changes in the fraction of fura-2 that has a bound Ca2+ ion and thus in the concentration of cytosolic Ca2+ (Figure 5-8).

Figure 5-8. Changes in the local concentration of Ca2+in a sea urchin egg following fertilization.

Figure 5-8

Changes in the local concentration of Ca2+in a sea urchin egg following fertilization. The Ca2+ throughout the cell was monitored at different times after fertilization using a fluorescence microscope and fura-2, a Ca2+-binding dye whose fluorescence (more...)

The fluorescence of other dyes is sensitive to the H+ concentration and can be used in a similar way to monitor the cytosolic pH of living cells.

Confocal Scanning and Deconvolution Microscopy Provide Sharper Images of Three-Dimensional Objects

Immunofluorescence microscopy has its limitations. The fixatives employed to preserve cell architecture often destroy the antigenicity of a protein, that is, its ability to bind to its specific antibody. Also, the method generally gives poor results with thin cell sections, because embedding media often fluoresce themselves, obscuring the specific signal from the antibody. Moreover, in microscopy of whole cells, the fluorescent light comes from molecules above and below the plane of focus; thus the observer sees a superposition of fluorescent images from molecules at many depths in the cell, making it difficult to determine the actual three-dimensional molecular arrangement (see Figure 5-6).

The confocal scanning microscope avoids the last problem by permitting the observer to visualize fluorescent molecules in a single plane of focus, thereby creating a vastly sharper cross-sectional image (Figure 5-9). At any instant during confocal imaging, only a single small part of a sample is illuminated with exciting light from a focused laser beam, which rapidly moves to different spots in the sample focal plane. Images from these spots are recorded by a video camera and stored in a computer, and the composite image is displayed on a computer screen.

Figure 5-9. The advantage of confocal fluorescence microscopy.

Figure 5-9

The advantage of confocal fluorescence microscopy. A mitotic fertilized egg from a sea urchin (Psammechinus) was lysed with a detergent, exposed to an anti-tubulin antibody, and then exposed to a fluorescein-tagged antibody that binds to the first antibody. (more...)

Deconvolution microscopy is similar to confocal microscopy in that a cross-sectional image is obtained, but the two techniques differ in the details of how this image is generated. In both cases, the objective lens collects light that originates from above and below the focal plane as well as that which originates from within the focal plane. Confocal microscopes use a pinhole to exclude the out-of-focus light. In contrast, deconvolution microscopes collect all the light from several focal planes, and then mathematically reassign the out-of-focus light to its correct focal plane with the aid of a high-speed computer, a mathematical operation called deconvolution.

To understand how a deconvolution microscope works, consider an infinitely small fluorescent source of light, which can be approximated by a fluorescent bead smaller than the resolution of the light microscope (i.e., <0.2 μm in diameter). The emitted light radiates in all directions, and when the source is in the focal plane of the objective, it appearsas a bright point of light. When the point source is outside the focal plane of the objective, some of the light is still collected by the objective lens, and the point source appears as a halo. As the focal plane is moved farther away from the plane containing the point source, the halo becomes larger and more diffuse. Knowing exactly how the light emitted by an infinitely small fluorescent source is collected and distorted by the optics of the sample and microscope, it is possible to reconstruct an individual cross-sectional image (containing only light that originated in the focal plane of interest) from a set of images taken as the objective focal plane is moved through the plane of interest.

Cross-sectional images obtained with a deconvolution microscope may have even greater detail than those obtained with a confocal microscope. Additionally, the fluorescent labeling of the sample does not need to be as intense for deconvolution microscopy as it does for confocal microscopy, since all the light produced by a fluorescent sample is collected and analyzed by the microscope. Three-dimensional images can be obtained by a refinement known as optical sectioning. In this method, a computer records individual fluorescent images of planes at different depths of the sample — in effect, serial sections — and combines the stack of images into one three-dimensional image (Figure 5-10).

Figure 5-10. Optical sectioning of a developing Drosophila egg chamber obtained with deconvolution fluorescence microscopy.

Figure 5-10

Optical sectioning of a developing Drosophila egg chamber obtained with deconvolution fluorescence microscopy. An egg chamber was labeled with the dye DAPI, which binds to DNA and generates a blue fluorescence from the nuclei. Actin filaments were labeled (more...)

Phase-Contrast and Nomarski Interference Microscopy Visualize Unstained Living Cells

Detailed views of transparent, live, unstained cells and tissues are obtainable with phase-contrast microscopy and Nomarski interference microscopy. Both techniques take advantage of the phenomena of refraction and diffraction of light waves (Figure 5-11). As a result, small differences in refractive index and thickness between parts of the specimen (say, between the nucleus and cytosol) or between the specimen and the surrounding medium can be converted into differences of light and dark in the final image.

Figure 5-11. Light passing through a specimen can be redirected by refraction and diffraction.

Figure 5-11

Light passing through a specimen can be redirected by refraction and diffraction. (a) Refraction: Because light moves at different speeds in different materials (more slowly in a medium of higher refractive index), a beam of light is bent (refracted) (more...)

The phase-contrast microscope generates an image in which the degree of darkness or brightness of a region of the sample depends on the refractive index of that region (Figure 5-12). The improved definition of subcellular structures in live, unstained cells obtained by phase-contrast microscopy compared with standard bright-field microscopy is illustrated in Figure 5-13a.

Figure 5-12. The optical pathway of the phase-contrast microscope.

Figure 5-12

The optical pathway of the phase-contrast microscope. Incident light passes through an annular diaphragm, and the condenser lens focuses a circular annulus (ring) of light on the sample. Light that passes unobstructed through the specimen is focused by (more...)

Figure 5-13. Phase contrast and Nomarski optics.

Figure 5-13

Phase contrast and Nomarski optics. (a) Several live, cultured fat cells (adipocytes) viewed by bright-field microscopy (left) and phase-contrast microscopy (right). (b) Another specimen of adipocytes viewed with Nomarski interference (differential interference) (more...)

Nomarski, or differential, interference microscopy generates an image that looks as if the specimen is casting a shadow to one side (Figure 5-13b): the “shadow” primarily represents a difference in refractive index and thickness of a specimen rather than its topography. In this technique, a prism splits an incident beam of plane-polarized light so that one part of the beam passes through one region of a specimen and the other part passes through a closely adjacent region; a second prism then reassembles the two beams. Minute differences in thickness or in the refractive index between adjacent parts of a sample are converted into a bright image (if the two beams are in phase when they recombine) or a dark one (if they are out of phase).

Phase-contrast microscopy is especially useful in examining the structure and movement of larger organelles, such as the nucleus and mitochondria, in live cultured cells. The greatest disadvantage of this technique is that it is suitable for observing only single cells or thin cell layers. Nomarski interference microscopy, in contrast, defines only the outlines of large organelles, such as the nucleus and vacuole. However, thick objects, such as the nuclei in a worm, can be observed by combining this technique with optical sectioning (Figure 5-13c).

Both phase-contrast and Nomarski interference microscopy can be used in time-lapse microscopy, in which the same cell is photographed at regular intervals over periods of several hours. This procedure allows the observer to study cell movement, provided the microscope’s stage can control the temperature of the specimen and the gas environment (Figure 5-14).

Figure 5-14. Time-lapse micrographs show the movement of a cultured fibroblast cell along a glass surface.

Figure 5-14

Time-lapse micrographs show the movement of a cultured fibroblast cell along a glass surface. A bit of debris on the substratum serves as a reference point. The first image, at 0 min, was obtained by phase-contrast microscopy. Successive images of the (more...)

Transmission Electron Microscopy Has a Limit of Resolution of 0.1 nm

The fundamental principles of electron microscopy are similar to those of light microscopy; the major difference is that electromagnetic lenses, not optical lenses, focus a high velocity electron beam instead of visible light. Because electrons are absorbed by atoms in air, the entire tube between the electron source and the viewing screen is maintained under an ultrahigh vacuum.

The transmission electron microscope (TEM) directs a beam of electrons through a specimen. Electrons are emitted by a tungsten cathode when it is electrically heated. The electric potential of the cathode is kept at 50,000 – 100,000 volts; that of the anode, near the top of the tube, is zero. This drop in voltage causes the electrons to accelerate as they move toward the anode. A condenser lens focuses the electron beam onto the sample; objective and projector lenses focus the electrons that pass through the specimen and project them onto a viewing screen or a piece of photographic film (Figure 5-15).

Figure 5-15. The optical path in a transmission electron microscope.

Figure 5-15

The optical path in a transmission electron microscope. A beam of electrons emanating from a heated tungsten filament is focused onto the specimen plane by the magnetic condenser lens. The electrons passing through the specimen are focused by a series (more...)

In typical electron microscopes, electrons have the properties of a wave with a wavelength of only 0.005 nm. Recall that the minimum distance D at which two objects can be distinguished is proportional to the wavelength λ of the light that illuminates the objects. Thus the limit of resolution for the electron microscope is theoretically 0.005 nm (less than the diameter of a single atom), or 40,000 times better than the resolution of the light microscope and 2 million times better than that of the unaided human eye. However, the effective resolution of the electron microscope in the study of biological systems is considerably less than this ideal. Under optimal conditions, a resolution of 0.10 nm can be obtained with transmission electron microscopes, about 2000 times better than the best resolution of light microscopes.

Preparation of Fixed, Stained Samples for TEM

Like the light microscope, the transmission electron microscope is used to view thin sections of a specimen, but the fixed sections must be much thinner for electron microscopy (only 50 – 100 nm, about 0.2 percent of the thickness of a single cell). Clearly, only a small portion of a cell can be observed in any one section. Figure 5-16 depicts the preparation of a sample for transmission electron microscopy. Generation of the image depends on differential scattering of the incident electrons by molecules in the preparation. Without staining, the beam of electrons passes through a cell or tissue sample uniformly, so the entire sample appears uniformly bright with little differentiation of components. Staining techniques are therefore used to reveal the location and distribution of specific materials.

Figure 5-16. Preparation of a sample of tissue for transmission electron microscopy.

Figure 5-16

Preparation of a sample of tissue for transmission electron microscopy. The tissue is dissected, cut into small cubes, and plunged into a fixing solution that cross-links and immobilizes proteins. (Glutaraldehyde is frequently used; osmium tetroxide, (more...)

Heavy metals, such as gold or osmium, appear dark on a micrograph because they scatter (diffract) most of the incident electrons; scattered electrons are not focused by the electromagnetic lenses and do not form the image. Osmium tetrox-ide preferentially stains certain cellular components, such as membranes, which appear black in micrographs. Specific proteins can be detected in thin sections by use of electron-dense gold particles coated with protein A, a bacterial protein that binds antibody molecules nonspecifically (Figure 5-17).

Figure 5-17. Localization of a specific protein by electron microscopy of antibody-stained samples.

Figure 5-17

Localization of a specific protein by electron microscopy of antibody-stained samples. (a) First the antibody is allowed to interact with its specific antigen (e.g., catalase) in a section of fixed tissue. Then the section is treated with a complex of (more...)

Electron microscopy also is used to obtain information about the shapes of purified viruses, fibers, enzymes, and other subcellular particles. In one technique, called metal shadowing, a thin layer of evaporated metal, such as platinum, is laid at an angle on a biological sample (Figure 5-18). An acid bath dissolves the biological material, leaving a metal replica of its surface, which can then be examined in the transmission electron microscope. Variations in the angle and thickness of the deposited metal allow an image to be formed because some incident electrons will be scattered in various directions rather than pass through the preparation. If the metal is deposited mainly on one side of the sample, for instance, the image seems to have “shadows,” where the metal appears dark and the shadows appear light.

Figure 5-18. Metal shadowing, a technique that makes surface details on very small particles visible in the electron microscope.

Figure 5-18

Metal shadowing, a technique that makes surface details on very small particles visible in the electron microscope. The sample is spread on a mica surface and then dried in a vacuum evaporator. A filament of a heavy metal, such as platinum or gold, is (more...)

Cryoelectron Microscopy

Standard electron microscopy cannot be used to study live cells because they are generally too vulnerable to the required conditions and preparatory techniques. In particular, the absence of water causes macromolecules to become denatured and nonfunctional. However, the technique of cryoelectron microscopy allows examination of hydrated, unfixed, and unstained biological specimens directly in the transmission electron microscope. In this technique, an aqueous suspension of a sample is applied in an extremely thin film to a grid. After it has been frozen in liquid nitrogen and maintained in this state by means of a special mount, it is observed in the electron microscope. The very low temperature (−196 °C) keeps the water from evaporating, even in a vacuum, and the sample can be observed in detail in its native, hydrated state without shadowing or fixing it (Figure 5-19). By computer-based averaging of images of hundreds of particles, a three-dimensional model almost to atomic resolution can be generated.

Figure 5-19. Cryoelectron micrograph of unstained rotavirus particles.

Figure 5-19

Cryoelectron micrograph of unstained rotavirus particles. A thin suspension of virus particles in water is applied to an electron microscopy grid and frozen. It is then visualized in a transmission electron microscope equipped with a sample stage cooled (more...)

Scanning Electron Microscopy Visualizes Details on the Surfaces of Cells and Particles

The scanning electron microscope allows the investigator to view the surfaces of unsectioned specimens. These cannot be visualized with transmission equipment because the electrons pass through the entire specimen. The sample is fixed, dried, and coated with a thin layer of a heavy metal, such as platinum, by evaporation in a vacuum (see Figure 5-18); in this case, the sample is rotated so that the platinum is deposited uniformly on the surface. An intense electron beam inside the microscope scans rapidly over the sample. Molecules in the specimen are excited and release secondary electrons that are focused onto a scintillation detector; the resulting signal is displayed on a cathode-ray tube. Because the number of secondary electrons produced by any one point on the sample depends on the angle of the electron beam in relation to the surface, the scanning electron micrograph has a threedimensional appearance (Figure 5-20; see also Figure 5-1a). The resolving power of scanning electron microscopes, which is limited by the thickness of the metal coating, is only about 10 nm, much less than that of transmission instruments.

Figure 5-20. Scanning electron micrograph of a tendon located within the shoulder of a 29-year-old male.

Figure 5-20

Scanning electron micrograph of a tendon located within the shoulder of a 29-year-old male. The sample was rapidly frozen and metal-shadowed before viewing in a scanning electron microscope. These long collagen fibrils are of variable diameter and are (more...)


  •  Various microscopic techniques generate different views of the cell and have different resolutions. The limit of resolution of a light microscope is about 0.2 μm; of a transmission electron microscope, about 0.1 nm; and of a scanning electron microscope, about 10 nm.
  •  Standard (bright-field) light microscopy is best for stained or colored cells or tissue sections.
  •  Fluorescence microscopy allows specific proteins and organelles to be detected in fixed cells stained with a fluorescent dye or fluorescent-labeled antibodies (immunofluorescence microscopy). The movements of microinjected or expressed recombinant fluorescent proteins also can be followed in living cells.
  •  By use of dyes whose fluorescence is proportional to the concentration of Ca2+ or H+ ions, fluorescence microscopy can measure the local concentration of Ca2+ ions and intracellular pH in living cells.
  •  Confocal imaging, which allows the observer to view fluorescent molecules in a single plane of a specimen, permits optical sectioning of the sample and produces very sharp images.
  •  Phase-contrast and Nomarski optics enable scientists to view the details of live, unstained cells and to monitor cell movement.
  •  Specimens for electron microscopy generally must be fixed, sectioned, and dehydrated, and then stained with electron-dense heavy metals.
  •  Surface details of particles such as viruses and collagen fibers can be revealed by electron microscopy of metal-shadowed specimens (see Figure 5-18).
  •  Unfixed, unstained specimens can be viewed in the electron microscope if they are frozen in hydrated form, a technique called cryoelectron microscopy.
  •  The scanning electron microscope can be used to view unsectioned cells or tissues; it produces images that appear to be three-dimensional.

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