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Cell_Biology_Alberts. Figure 9–37 Superresolution microscopy can be achieved by reducing the size of the point spread function. (a) the size of a normal focused beam of excitatory light. (B) an extremely strong superimposed laser beam, at a different wavelength and in the shape of a torus, depletes emitted fluorescence everywhere in the specimen except right in the center of the beam, reducing the effective width of the point spread function (C). as the specimen is scanned, this small point spread function can then build up a crisp image in a process called steD (stimulated emission depletion microscopy). (D) synaptic vesicles in live cultured neurons, fluorescently labeled and imaged by ordinary confocal microscopy, with a resolution of 260 nm. (e) the same vesicles imaged by steD, with a resolution of 60 nm, which allows single vesicles to be resolved. (F) Fluorescently labeled replication factories in the nucleus of a cultured cell, imaged by ordinary confocal microscopy. (g) the same replication


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Cell_Biology_Alberts. 60 nm, which allows single vesicles to be resolved. (F) Fluorescently labeled replication factories in the nucleus of a cultured cell, imaged by ordinary confocal microscopy. (g) the same replication factories imaged by steD. single, discrete replication sites can be resolved by steD that cannot be seen in the confocal image. (a, B, and C, from g. Donnert et al., Proc. Natl Acad. Sci. USA 103:11440–11445, 2006. With permission from national academy of sciences; D and e, from V. Westphal et al., Science 320:246– 249, 2008. With permission from aaas; F and g, from Z. Cseresnyes,


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Cell_Biology_Alberts. U. schwarz and C.M. green, BMC Cell Biol. 10:88, 2009.) of adjacent fluorescent molecules, as we saw earlier, is that they each contribute blurry, overlapping point spread functions to the image, making the exact position of any one molecule impossible to resolve. Another way round this limitation is to arrange for only a very few, clearly separated molecules to actively fluoresce at any one moment. The exact position of each of these can then be computed, before subsequent sets of molecules are examined.


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Cell_Biology_Alberts. In practice, this can be achieved by using lasers to sequentially switch on a sparse subset of fluorescent molecules in a specimen containing photoactivatable or photoswitchable fluorescent labels. Labels are activated, for example, by illumination with nearultraviolet light, which modifies a small subset of molecules so that they fluoresce when exposed to an excitation beam at another wavelength. These are then imaged before bleaching quenches their fluorescence and a new subset is activated. Each molecule emits a few thousand photons in response to the excitation before switching off, and the switching process can be repeated hundreds or even thousands of times, allowing the exact coordinates of a very large set of single molecules to be determined. The full set can be combined and digitally displayed as an image in which the computed location of each individual molecule is exactly marked (Figure 9–38). This class of methods has been variously termed photoactivated localization


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Cell_Biology_Alberts. digitally displayed as an image in which the computed location of each individual molecule is exactly marked (Figure 9–38). This class of methods has been variously termed photoactivated localization microscopy (PALM) or stochastic optical reconstruction microscopy (STORM).


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Cell_Biology_Alberts. By switching the fluorophores off and on sequentially in different regions of the specimen as a function of time, all the superresolution imaging methods described above allow the resolution of molecules that are much closer together than the 200 nm diffraction limit. In STED, the locations of the molecules are determined by using optical methods to define exactly where their fluorescence will be on or off. In PALM and STORM, individual fluorescent molecules are switched on and off at random over a period of time, allowing their positions to be accurately determined. PALM and STORM techniques have depended on the 100 photons 1000 photons 10,000 photons successive cycles of activation and bleaching allow well-separated single fuorescent molecules to be detected the exact center of each fuorescent molecule is determined and its position added to the map (C)a super-resolution image of the fuorescent structure is built up as the positions of successive small 1 µmgroups of molecules are


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Cell_Biology_Alberts. molecule is determined and its position added to the map (C)a super-resolution image of the fuorescent structure is built up as the positions of successive small 1 µmgroups of molecules are added to the map


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Cell_Biology_Alberts. Figure 9–38 Single fluorescent molecules can be located with great accuracy. (a) Determining the exact mathematical center of the blurred image of a single fluorescent molecule becomes more accurate the more photons contribute to the final image. the point spread function described in the text dictates that the size of the molecular image is about 200 nm across, but in very bright specimens, the position of its center can be pinpointed to within a nanometer. (B) in this imaginary specimen, sparse subsets of fluorescent molecules are individually switched on briefly and then bleached. the exact positions of all these well-spaced molecules can be gradually built up into an image at superresolution. (C) in this portion of a cell, the microtubules have been fluorescently labeled and imaged at the top in a tirF microscope (see Figure 9–32) and below, at superresolution, in a paLM microscope. the diameter of the microtubules in the lower panel now resembles their true size, about 25 nm,


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Cell_Biology_Alberts. the top in a tirF microscope (see Figure 9–32) and below, at superresolution, in a paLM microscope. the diameter of the microtubules in the lower panel now resembles their true size, about 25 nm, rather than the 250 nm in the blurred image at the top. (a, from a.L. Mcevoy et al., BMC Biol. 8:106, 2010; C, courtesy of Carl Zeiss Ltd.) development of novel fluorescent probes that exhibit the appropriate switching behavior. All these methods are now being extended to incorporate multicolor imaging, threedimensional imaging (Figure 9–39), and livecell imaging in real time. Ending the long reign of the diffraction limit has certainly reinvigorated light microscopy and its place in cell biology research.


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Cell_Biology_Alberts. Many light-microscope techniques are available for observing cells. Cells that have been fixed and stained can be studied in a conventional light microscope, whereas antibodies coupled to fluorescent dyes can be used to locate specific molecules in cells in a fluorescence microscope. Living cells can be seen with phase-contrast, differential-interference-contrast, dark-field, or bright-field microscopes. All forms of light microscopy are facilitated by digital image-processing techniques, which enhance sensitivity and refine the image. Confocal microscopy and image deconvolution both provide thin optical sections and can be used to reconstruct three-dimensional images.


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Cell_Biology_Alberts. Techniques are now available for detecting, measuring, and following almost any desired molecule in a living cell. Fluorescent indicator dyes can be introduced to measure the concentrations of specific ions in individual cells or in different parts of a cell. Virtually any protein of interest can be genetically engineered as a fluorescent fusion protein, and then imaged in living cells by fluorescence microscopy. The dynamic behavior and interactions of many molecules can be followed in living cells by variations on the use of fluorescent protein tags, in some cases at the level of single molecules. Various superresolution techniques can circumvent the diffraction limit and resolve molecules separated by distances as small as 20 nm. Looking at CeLLs anD MoLeCULes in the eLeCtron MiCrosCope


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Cell_Biology_Alberts. Looking at CeLLs anD MoLeCULes in the eLeCtron MiCrosCope Light microscopy is limited in the fineness of detail that it can reveal. Microscopes using other types of radiation—in particular, electron microscopes—can resolve much smaller structures than is possible with visible light. This higher resolution comes at a cost: specimen preparation for electron microscopy is complex and it is harder to be sure that what we see in the image corresponds precisely to the original living structure. It is possible, however, to use very rapid freezing to preserve structures faithfully for electron microscopy. Digital image analysis can be used to reconstruct threedimensional objects by combining information either from many individual particles or from multiple tilted views of a single object. Together, these approaches extend the resolution and scope of electron microscopy to the point at which we can faithfully image the structures of individual macromolecules and the complexes they form.


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Cell_Biology_Alberts. the electron Microscope resolves the Fine structure of the Cell


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Cell_Biology_Alberts. The formal relationship between the diffraction limit to resolution and the wavelength of the illuminating radiation (see Figure 9–6) holds true for any form of radiation, whether it is a beam of light or a beam of electrons. With electrons, however, the limit of resolution is very small. The wavelength of an electron decreases as its velocity increases. In an electron microscope with an accelerating voltage of 100,000 V, the wavelength of an electron is 0.004 nm. In theory, the resolution of such a microscope should be about 0.002 nm, which is 100,000 times that of the light microscope. Because the aberrations of an electron lens are considerably harder to correct than those of a glass lens, however, the practical resolving power of modern electron microscopes is, even with careful image processing to correct for lens aberrations, about 0.05 nm (0.5 Å) (Figure 9–40). This is because only the very center of the electron lenses can be used, and the effective numerical aperture is tiny.


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Cell_Biology_Alberts. processing to correct for lens aberrations, about 0.05 nm (0.5 Å) (Figure 9–40). This is because only the very center of the electron lenses can be used, and the effective numerical aperture is tiny. Furthermore, problems of specimen preparation, contrast, and radiation damage have generally limited the normal effective resolution for biological objects to 1 nm (10 Å). This is nonetheless about 200 times better than the resolution of the light microscope. Moreover, the performance of electron microscopes is improved by electron illumination sources called field emission guns. These very bright and coherent sources substantially improve the resolution achieved.


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Cell_Biology_Alberts. In overall design, the transmission electron microscope (TEM) is similar to a light microscope, although it is much larger and “upside down” (Figure 9–41). 0.14 nm Figure 9–40 The resolution of the electron microscope. this transmission electron micrograph of a monolayer of graphene resolves the individual carbon atoms as bright spots in a hexagonal lattice. graphene is a single isolated atomic plane of graphite and forms the basis of carbon nanotubes. the distance between adjacent bonded carbon atoms is 0.14 nm (1.4 Å). such resolution can only be obtained in a specially built transmission electron microscope in which all lens aberrations are carefully corrected, and with optimal specimens; it cannot be achieved with most conventional biological specimens. (From a. Dato et al., Chem. Commun. 40:6095–6097, 2009. With permission from the royal society of Chemistry.)


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Cell_Biology_Alberts. The source of illumination is a filament or cathode that emits electrons at the top of a cylindrical column about 2 m high. Since electrons are scattered by collisions with air molecules, air must first be pumped out of the column to create a vacuum. The electrons are then accelerated from the filament by a nearby anode and allowed to pass through a tiny hole to form an electron beam that travels down the column. Magnetic coils placed at intervals along the column focus the electron beam, just as glass lenses focus the light in a light microscope. The specimen is put into the vacuum, through an airlock, into the path of the electron beam. As in light microscopy, the specimen is usually stained—in this case, with electron-dense material. Some of the electrons passing through the specimen are scattered by structures stained with the electrondense material; the remainder are focused to form an image, in a manner analogous to the way an image is formed in a light microscope. The image can


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Cell_Biology_Alberts. scattered by structures stained with the electrondense material; the remainder are focused to form an image, in a manner analogous to the way an image is formed in a light microscope. The image can be observed on a phosphorescent screen or recorded with a highresolution digital camera. Because the scattered electrons are lost from the beam, the dense regions of the specimen show up in the image as areas of reduced electron flux, which look dark.


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Cell_Biology_Alberts. In the early days of its application to biological materials, the electron microscope revealed many previously unimagined structures in cells. But before these discoveries could be made, electron microscopists had to develop new procedures for embedding, cutting, and staining tissues.


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Cell_Biology_Alberts. Since the specimen is exposed to a very high vacuum in the electron microscope, living tissue is usually killed and preserved by fixation—first with glutaraldehyde, which covalently crosslinks protein molecules to their neighbors, and then with osmium tetroxide, which binds to and stabilizes lipid bilayers as well as proteins (Figure 9–42). Because electrons have very limited penetrating power, the fixed tissues normally have to be cut into extremely thin sections (25–100 nm thick, about 1/200 the thickness of a single cell) before they are viewed. This is achieved by dehydrating the specimen, permeating it with a monomeric resin that polymerizes to form a solid block of plastic, then cutting the block with a fine glass or diamond knife on a special microtome. The resulting thin sections, free of water and other volatile solvents, are supported on a small metal grid for viewing in the microscope (Figure 9–43).


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Cell_Biology_Alberts. Figure 9–41 The principal features of a light microscope and a transmission electron microscope. these drawings emphasize the similarities of overall design. Whereas the lenses in the light microscope are made of glass, those in the electron microscope are magnetic coils. the electron microscope requires that the specimen be placed in a vacuum. the inset shows a transmission electron microscope in use. (photograph courtesy of JeoL Ltd.) Figure 9–42 Two common chemical fixatives used for electron microscopy. the two reactive aldehyde groups of glutaraldehyde enable it to cross-link various types of molecules, forming covalent bonds between them. osmium tetroxide forms cross-linked complexes with many organic compounds, and in the process becomes reduced. this reaction is especially useful for fixing cell membranes, since the C=C double bonds present in many fatty acids react with osmium tetroxide.


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Cell_Biology_Alberts. The steps required to prepare biological material for electron microscopy are challenging. How can we be sure that the image of the fixed, dehydrated, resinembedded specimen bears any relation to the delicate, aqueous biological system present in the living cell? The best current approaches to this problem depend on rapid freezing. If an aqueous system is cooled fast enough and to a low enough temperature, the water and other components in it do not have time to rearrange themselves or crystallize into ice. Instead, the water is supercooled into a rigid but noncrystalline state—a “glass”—called vitreous ice. This state can be achieved by slamming the specimen onto a polished copper block cooled by liquid helium, by plunging it into or spraying it with a jet of a coolant such as liquid propane, or by cooling it at high pressure.


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Cell_Biology_Alberts. Some rapidly frozen specimens can be examined directly in the electron microscope using a special cooled specimen holder. In other cases, the frozen block can be fractured to reveal interior cell surfaces, or the surrounding ice can be sublimed away to expose external surfaces. However, we often want to examine thin sections. A compromise is therefore to rapidfreeze the tissue, replace the water with organic solvents, embed the tissue in plastic resin, and finally cut sections and stain. Although technically still difficult, this approach stabilizes and preserves the tissue in a condition very close to its original living state (Figure 9–44).


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Cell_Biology_Alberts. Image clarity in an electron micrograph depends upon having a range of contrasting electron densities within the specimen. Electron density in turn depends on the atomic number of the atoms that are present: the higher the atomic number, the more electrons are scattered and the darker that part of the image. Biological tissues are composed mainly of atoms of very low atomic number (primarily carbon, oxygen, nitrogen, and hydrogen). To make them visible, tissues are usually impregnated (before or after sectioning) with the salts of heavy metals such as uranium, lead, and osmium. The degree of impregnation, or “staining,” with these salts will vary for different cell constituents. Lipids, for example, tend to stain darkly after osmium fixation, revealing the location of cell membranes. We have seen how antibodies can be used in conjunction with fluorescence microscopy to localize specific macromolecules. An analogous method—immunogold copper grid covered with carbon and/or plastic flm


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Cell_Biology_Alberts. Figure 9–43 The metal grid that supports the thin sections of a specimen in a transmission electron microscope. Figure 9–44 Thin section of a cell. this thin section is of a yeast cell that has been very rapidly frozen and the vitreous ice replaced by organic solvents and then by plastic resin. the nucleus, mitochondria, cell wall, golgi stacks, and ribosomes can all be readily seen in a state that is presumed to be as lifelike as possible. (Courtesy of andrew staehelin.) 0.5 µm electron microscopy—can be used in the electron microscope. The usual procedure is to incubate a thin section first with a specific primary antibody, and then with a secondary antibody to which a colloidal gold particle has been attached. The gold particle is electrondense and can be seen as a black dot in the electron microscope (Figure 9–45). Different antibodies can be conjugated to different sized gold particles so multiple proteins can be localized in a single sample.


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Cell_Biology_Alberts. A complication for immunogold labeling is that the antibodies and colloidal gold particles do not penetrate into the resin used for embedding; therefore, they detect antigens only at the surface of the section. This means that the method’s sensitivity is low, since antigen molecules in the deeper parts of the section are not detected. Furthermore, we may get a false impression regarding which structures contain the antigen and which do not. One solution is to label the specimen before embedding it in plastic, when cells and tissues are still fully accessible to labeling reagents. Extremely small gold particles, about 1 nm in diameter, work best for this procedure. Such small gold particles are usually not easily visible in the final sections, so additional silver or gold is nucleated around the tiny 1 nm gold particles in a chemical process very much like photographic development. Different Views of a single object Can Be Combined to give a three-Dimensional reconstruction


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Cell_Biology_Alberts. Different Views of a single object Can Be Combined to give a three-Dimensional reconstruction Thin sections often fail to convey the threedimensional arrangement of cellular components viewed in a TEM, and the image can be very misleading: a linear structure such as a microtubule may appear in section as a pointlike object, for example, and a section through protruding parts of a single irregularly shaped solid body may give the appearance of two or more separate objects (Figure 9–46). The third dimension can be reconstructed from serial sections, but this is a lengthy and tedious process. Even thin sections, however, have a significant depth compared with the resolution of the electron microscope, so the TEM image can also be misleading in an opposite way, through the superimposition of objects that lie at different depths.


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Cell_Biology_Alberts. Figure 9–45 Localizing proteins in electron microscopy. immunogold electron microscopy is used here to find the specific location of four different protein components within the spindle pole body of yeast. at the top is a thin section of a yeast mitotic spindle showing the spindle microtubules that cross the nucleus and connect at each end to spindle pole bodies embedded in the nuclear envelope. a diagram of the components of a single spindle pole body is shown below. on separate sections, antibodies against four different proteins of the spindle pole body are used, together with colloidal gold particles (black dots), to reveal where within the complex structure each protein is located. (Courtesy of John kilmartin.)


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Cell_Biology_Alberts. Because of the large depth of field of electron microscopes, all the parts of the threedimensional specimen are in focus, and the resulting image is a projection (a superimposition of layers) of the structure along the viewing direction. The lost information in the third dimension can be recovered if we have views of the same specimen from many different directions. The computational methods for this technique are widely used in medical CT scans. In a CT scan, the imaging equipment is moved around the patient to generate the different views. In electron-microscope (EM) tomography, the specimen holder is tilted in the microscope, which achieves the same result. In this way, we can arrive at a threedimensional reconstruction, in a chosen standard orientation, by combining different views of a single object. Each individual view will be very noisy but by combining them in three dimensions and taking an average, the noise can be largely eliminated. Starting with thick plastic sections of


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Cell_Biology_Alberts. a single object. Each individual view will be very noisy but by combining them in three dimensions and taking an average, the noise can be largely eliminated. Starting with thick plastic sections of embedded material, threedimensional reconstructions, or tomograms, are used extensively to describe the detailed anatomy of specific regions of the cell, such as the Golgi apparatus (Figure 9–47) or the cytoskeleton. Increasingly, microscopists are also applying EM tomography to unstained frozen, hydrated sections, and even to rapidly frozen whole cells or organelles (Figure 9–48). Electron microscopy now provides a robust bridge between the scale of the single molecule and that of the whole cell.


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Cell_Biology_Alberts. images of surfaces Can Be obtained by scanning electron Microscopy A scanning electron microscope (SEM) directly produces an image of the threedimensional structure of the surface of a specimen. The SEM is usually smaller, simpler, and cheaper than a transmission electron microscope. Whereas the TEM uses the electrons that have passed through the specimen to form an Figure 9–46 A three-dimensional reconstruction from serial sections. single thin sections in the electron microscope sometimes give misleading impressions. in this example, most sections through a cell containing a branched mitochondrion seem to contain two or three separate mitochondria (compare Figure 9–44). sections 4 and 7, moreover, might be interpreted as showing a mitochondrion in the process of dividing. the true three-dimensional shape can be reconstructed from a complete set of serial sections.


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Cell_Biology_Alberts. Figure 9–47 Electron-microscope (EM) tomography. samples that have been rapidly frozen, and then freeze-substituted and embedded in plastic, preserve their structure in a condition that is very close to their original living state (Movie 9.2). this example shows the three-dimensional structure of the golgi apparatus from a rat kidney cell. several thick sections (250 nm) of the cell were tilted in a high-voltage electron microscope, along two different axes, and about 160 different views recorded. the digital data allow individual thin slices of the complete three-dimensional data set, or tomogram, to be viewed; for example, the serial slices, each only 4 nm thick, are shown in (a) and (B). Very little changes from one slice to the next, but using the full data set, and manually color-coding the membranes (B), one can obtain a full three-dimensional reconstruction, at a resolution of about 7 nm, of the complete golgi complex and its associated vesicles (C). (From M.s. Ladinsky et


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Cell_Biology_Alberts. the membranes (B), one can obtain a full three-dimensional reconstruction, at a resolution of about 7 nm, of the complete golgi complex and its associated vesicles (C). (From M.s. Ladinsky et al., J.Cell Biol. 144:1135–1149, 1999. With permission from the authors.)


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Cell_Biology_Alberts. Looking at CeLLs anD MoLeCULes in the eLeCtron MiCrosCope


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Cell_Biology_Alberts. Figure 9–48 Combining cryoelectronmicroscope tomography and single-particle reconstruction. small, unfixed, rapidly frozen specimens can be examined while still frozen. in this example, the small nuclei of the amoeba Dictyostelium were gently isolated and then very rapidly frozen before a series of angled views were recorded with the aid of a tilting microscope stage. these digital views are combined by eM tomographs to produce a three-dimensional tomogram. two thin digital slices (10 nm) through this tomogram show (a) top views and (B) side views of individual nuclear pores (white arrows). (C) in the three-dimensional model, a surface rendering of the pores (blue) is seen embedded in the nuclear envelope (yellow). From a series of tomograms it was possible to extract data sets for nearly 300 separate nuclear pores, whose structures could then be averaged using the techniques of single-particle reconstruction. the surface-rendered view of one of these reconstructed pores is shown (D)


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Cell_Biology_Alberts. 300 separate nuclear pores, whose structures could then be averaged using the techniques of single-particle reconstruction. the surface-rendered view of one of these reconstructed pores is shown (D) from the nuclear face and (e) in (E) 50 nm cross section (compare with Figure 12–8). the pore complex is colored blue and the nuclear basket brown. (From M. Beck image, the SEM uses electrons that are scattered or emitted from the specimen’s et al., Science 306:1387–1390, 2004. With permission from aaas.) surface. The specimen to be examined is fixed, dried, and coated with a thin layer of heavy metal. Alternatively, it can be rapidly frozen, and then transferred to a cooled specimen stage for direct examination in the microscope. Often an entire plant part or small animal can be put into the microscope with very little preparation (Figure 9–49). The specimen is scanned with a very narrow beam of electrons. The quantity of electrons scattered or emitted as this primary beam bombards each


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Cell_Biology_Alberts. microscope with very little preparation (Figure 9–49). The specimen is scanned with a very narrow beam of electrons. The quantity of electrons scattered or emitted as this primary beam bombards each successive point of the metallic surface is measured and used to control the intensity of a second beam, which moves in synchrony with the primary beam and forms an image on a computer screen. Eventually a highly enlarged image of the surface as a whole is built up (Figure 9–50).


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Cell_Biology_Alberts. The SEM technique provides great depth of field; moreover, since the amount of electron scattering depends on the angle of the surface relative to the beam, the image has highlights and shadows that give it a threedimensional appearance (see Figure 9–49 and Figure 9–51). Only surface features can be examined, however, and in most forms of SEM, the resolution attainable is not very high (about 10 nm, with an effective magnification of up to 20,000 times). As a result, the technique is usually used to study whole cells and tissues rather than subcellular organelles (see Movie 21.3). Veryhighresolution SEMs have, however, been developed with a bright coherentfield emission gun as the electron source. This type of SEM can produce images that rival the resolution possible with a TEM (Figure 9–52). negative staining and Cryoelectron Microscopy Both allow Macromolecules to Be Viewed at high resolution


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Cell_Biology_Alberts. If they are shadowed with a heavy metal to provide contrast, isolated macromolecules such as DNA or large proteins can be visualized readily in the electron microscope, but negative staining allows finer detail to be seen. In this technique, the molecules are supported on a thin film of carbon and mixed with a Figure 9–49 A developing wheat flower, solution of a heavymetal salt such as uranyl acetate. After the sample has dried, or spike. this delicate flower spike was rapidly frozen, coated with a thin metal film, a very thin film of metal salt covers the carbon film everywhere except where it and examined in the frozen state with an has been excluded by the presence of an adsorbed macromolecule. Because the seM. this low-magnification micrograph macromolecule allows electrons to pass through it much more readily than does demonstrates the large depth of focus of the surrounding heavymetal stain, a reverse or negative image of the molecule is an seM. (Courtesy of kim Findlay.)


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Cell_Biology_Alberts. through it much more readily than does demonstrates the large depth of focus of the surrounding heavymetal stain, a reverse or negative image of the molecule is an seM. (Courtesy of kim Findlay.) created. Negative staining is especially useful for viewing large macromolecular aggregates such as viruses or ribosomes, and for seeing the subunit structure of protein filaments (Figure 9–53).


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Cell_Biology_Alberts. Shadowing and negative staining can provide highcontrast surface views of small macromolecular assemblies, but the size of the smallest metal particles in the shadow or stain used limits the resolution of both techniques. An alternative that allows us to visualize directly at high resolution even the interior features of threedimensional structures such as viruses and organelles is cryoelectron microscopy, in which rapid freezing to form vitreous ice is again the key. A very thin (about 100 nm) film of an aqueous suspension of virus or purified macromolecular complex is prepared on a microscope grid and is then rapidly frozen by


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Cell_Biology_Alberts. Figure 9–50 The scanning electron microscope. in an seM, the specimen is scanned by a beam of electrons brought to a focus on the specimen by the electromagnetic coils that act as lenses. the detector measures the quantity of electrons scattered or emitted as the beam bombards each successive point on the surface of the specimen and controls the intensity of successive points in an image built up on a screen. the seM creates striking images of three-dimensional objects with great depth of focus and a resolution between 3 nm and 20 nm depending on the instrument. (photograph courtesy of andrew Davies.)


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Cell_Biology_Alberts. Figure 9–51 Scanning electron microscopy. (a) a scanning electron micrograph of the stereocilia projecting from a hair cell in the inner ear of a bullfrog. For comparison, the same structure is shown by (B) differential-interferencecontrast light microscopy (Movie 9.3) and (C) thin-section transmission electron microscopy. (Courtesy of richard Jacobs and James hudspeth.) being plunged into a coolant. A special sample holder keeps this hydrated specimen at –160°C in the vacuum of the microscope, where it can be viewed directly without fixation, staining, or drying. Unlike negative staining, in which what we see is the envelope of stain exclusion around the particle, hydrated cryoelectron microscopy produces an image from the macromolecular structure itself. However, the contrast in this image is very low, and to extract the maximum amount of structural information, special imageprocessing techniques must be used, as we describe next.


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Cell_Biology_Alberts. Multiple images Can Be Combined to increase resolution As we saw earlier (p. 532), noise is important in light microscopy at low light levels, but it is a particularly severe problem for electron microscopy of unstained macromolecules. A protein molecule can tolerate a dose of only a few tens of electrons per square nanometer without damage, and this dose is orders of magnitude below what is needed to define an image at atomic resolution.


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Cell_Biology_Alberts. The solution is to obtain images of many identical molecules—perhaps tens of thousands of individual images—and combine them to produce an averaged image, revealing structural details that are hidden by the noise in the original images. This procedure is called single-particle reconstruction. Before combining all the individual images, however, they must be aligned with each other. Sometimes it is possible to induce proteins and complexes to form crystalline arrays, in which each molecule is held in the same orientation in a regular lattice. In this case, the alignment problem is easily solved, and several protein structures have been determined at atomic resolution by this type of electron crystallography. In principle, however, crystalline arrays are not absolutely required. With the help of a computer, the digital images of randomly distributed and unaligned molecules can be processed and combined to yield highresolution reconstructions (see Movie 13.1). Although structures that


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Cell_Biology_Alberts. help of a computer, the digital images of randomly distributed and unaligned molecules can be processed and combined to yield highresolution reconstructions (see Movie 13.1). Although structures that have some intrinsic symmetry make the task of alignment easier and more accurate, this technique has also been used for objects like ribosomes, with no symmetry. Figure 9–54 shows the structure of


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Cell_Biology_Alberts. Figure 9–52 The nuclear pore. rapidly frozen nuclear envelopes were imaged in a high-resolution seM, equipped with a field emission gun as the electron source. these views of each side of a nuclear pore represent the limit of resolution of the seM (compare with Figure 12–8). (Courtesy of Martin goldberg and terry allen.) We know in detail about many cell processes, such as Dna replication and transcription and rna translation, but will we ever be able to visualize such rapid molecular processes in action in cells? Will we ever be able to image intracellular structures at the resolution of the electron microscope in living cells? how can we improve crystallization and single-particle cryoelectron microscopy techniques to obtain high-resolution structures of all important membrane channels and transporters? What new concepts might these structures reveal?


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Cell_Biology_Alberts. Figure 9–53 Negatively stained actin filaments. in this transmission electron micrograph, each filament is about 8 nm in diameter and is seen, on close inspection, to be composed of a helical chain of globular actin molecules. (Courtesy of roger Craig.) the protein capsid inside a human immunodeficiency virus (HIV) that has been determined at high resolution by the combination of many particles, multiple views, and molecular modeling.


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Cell_Biology_Alberts. A resolution of 0.3 nm has been achieved by electron microscopy—enough to begin to see the internal atomic arrangements in a protein and to rival xray crystallography in resolution. Although electron microscopy is unlikely to supersede xray crystallography (discussed in Chapter 8) as a method for macromolecular structure determination, it has some very clear advantages. First, it does not absolutely require crystalline specimens. Second, it can deal with extremely large complexes—structures that may be too large or too variable to crystallize satisfactorily. Third, it allows the rapid analysis of different conformations of protein complexes.


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Cell_Biology_Alberts. The analysis of large and complex macromolecular structures is helped considerably if the atomic structure of one or more of the subunits is known, for example from xray crystallography. Molecular models can then be mathematically “fitted” into the envelope of the structure determined at lower resolution using the electron microscope (see Figures 16–16D and 16–46). Figure 9–55 shows the structure of a ribosome with the location of a bound release factor displayed in this way (see also Figure 6–72).


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Cell_Biology_Alberts. Discovering the detailed structure of membranes and organelles requires the higher resolution attainable in a transmission electron microscope. Specific macromolecules can be localized after being labeled with colloidal gold linked to antibodies. Three-dimensional views of the surfaces of cells and tissues are obtained by scanning electron microscopy. The shapes of isolated molecules can be readily determined by electron microscopy techniques involving fast freezing or negative staining. Electron tomography and single-particle reconstruction use computational manipulations of data obtained from multiple images and multiple viewing angles to produce detailed reconstructions of macromolecules and molecular complexes. The resolution obtained with these methods means that atomic structures of individual


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Cell_Biology_Alberts. Figure 9–54 Single-particle reconstruction. the structure of a complete human immunodeficiency virus (hiV) capsid has been determined by a combination of cryoelectron microscopy, protein structure determination, and modeling. (a) a single 4 nm slice from an eM tomographic model (see also Figure 9–48) of an intact hiV particle with its membrane outer envelope and its internal, irregularly shaped protein capsid that houses its rna genome. (B) electron microscopy of capsid subunits that have self-assembled into a helical tube can be used to derive an electron-density map at a resolution of 8 nm, in which details of the hexamers are clearly visible. (C) Using the known atomic coordinates of a single subunit of the hexamer, the structure has been modeled into the electron-density map from (B). (D) a molecular reconstruction of the entire hiV capsid, based on the detailed structures shown in (a) and (C). this capsid contains 216 hexamers (blue) and 12 pentamers (yellow). (adapted from g.


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Cell_Biology_Alberts. (D) a molecular reconstruction of the entire hiV capsid, based on the detailed structures shown in (a) and (C). this capsid contains 216 hexamers (blue) and 12 pentamers (yellow). (adapted from g. Zhao et al., Nature 497:643–646, 2013. With permission from Macmillan publishers Ltd. C, pDB code: 3J34.)


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Cell_Biology_Alberts. Figure 9–55 Single-particle reconstruction and molecular model fitting. Bacterial ribosomes, with and without the release factor required for peptide release from the ribosome, were used to derive high-resolution, three-dimensional cryoelectron microscopy maps at a resolution of better than 1 nm. images of nearly 20,000 separate ribosomes preserved in ice were used to produce single-particle reconstructions.


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Cell_Biology_Alberts. (a) the 30s ribosomal subunit (yellow) and the 50s subunit (blue) can be distinguished from the additional electron density that can be attributed to the release factor rF2 (purple). (B) the known molecular structure of rF2 modeled into the electron density from (a). (From U.B.s. rawat et al., Nature 421:87–90, 2003. With permission from Macmillan publishers Ltd.) macromolecules can often be “fitted” to the images derived by electron microscopy. In this way, the TEM is increasingly able to bridge the gap between structures discovered by x-ray crystallography and those discovered with the light microscope. Figure Q9–1 paths of light rays through dry and oil-immersion lenses (problem 9–3). the red circle at the origin of the light rays is the specimen. 9–4 Figure Q9–2 shows a diagram of the human eye. The refractive indices of the components in the light path are: cornea 1.38, aqueous humor 1.33, crystalline lens 1.41, and Which statements are true? Explain why or why not.


Cell_Biology_Alberts

Cell_Biology_Alberts. Which statements are true? Explain why or why not. 9–1 Because the DNA double helix is only 2 nm wide— well below the limit of resolution of the light microscope— it is impossible to see chromosomes in living cells without special stains. 9–2 A fluorescent molecule, having absorbed a single photon of light at one wavelength, always emits it at a longer wavelength. Discuss the following problems. 9–3 The diagrams in Figure Q9–1 show the paths of light rays passing through a specimen with a dry lens and with an oilimmersion lens. Offer an explanation for why oilimmersion lenses should give better resolution. Air, glass, and oil have refractive indices of 1.00, 1.51, and 1.51, 9–5 Why do humans see so poorly under water? And why do goggles help? 9–6 Explain the difference between resolution and magnification.


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Cell_Biology_Alberts. 9–6 Explain the difference between resolution and magnification. 9–7 Antibodies that bind to specific proteins are important tools for defining the locations of molecules in cells. The sensitivity of the primary antibody—the antibody that reacts with the target molecule—is often enhanced by using labeled secondary antibodies that bind to it. What are the advantages and disadvantages of using secondary antibodies that carry fluorescent tags versus those that carry bound enzymes? 9–8 Figure Q9–3 shows a series of modified fluorescent proteins that emit light in a range of colors. How do you suppose the exact same chromophore can fluoresce at so many different wavelengths?


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Cell_Biology_Alberts. Figure Q9–3 a rainbow of colors produced by modified fluorescent proteins (problem 9–8). (Courtesy of nathan shaner, paul steinbach and roger tsien.) 9–9 Consider a fluorescent detector designed to report the cellular location of active protein tyrosine kinases. A blue (cyan) fluorescent protein (CFP) and a yellow fluorescent protein (YFP) were fused to either end of a hybrid protein domain. The hybrid protein segment consisted of a substrate peptide recognized by the Abl protein tyrosine kinase and a phosphotyrosinebinding domain (Figure Q9–4A). Stimulation of the CFP domain does not cause emission by the YFP domain when the domains are 1.3 (problem 9–4). binding protein retina 1.0 vitreous humor 1.38. Where does the main refraction—the main focusing—occur? What role do you suppose the lens 1.2 plays? 1.1 iris of the human eye


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Cell_Biology_Alberts. 1.1 iris of the human eye Figure Q9–4 Fluorescent reporter protein designed to detect tyrosine phosphorylation (problem 9–9). (a) Domain structure of reporter protein. Four domains are indicated: CFp, YFp, tyrosine kinase substrate peptide, and a phosphotyrosine-binding domain. (B) Fret assay. YFp/CFp is normalized to 1.0 at time zero. the reporter was incubated in the presence (or absence) of abl and atp for the indicated times. Arrow indicates time of addition of a tyrosine phosphatase. (From a.Y. ting, k.h. kain, r.L. klemke and r.Y. tsien, Proc. Natl Acad. Sci. USA 98:15003–15008, 2001. With permission from national academy of sciences.) separated. When the CFP and YFP domains are brought close together, however, fluorescence resonance energy transfer (FRET) allows excitation of CFP to stimulate emission by YFP. FRET shows up experimentally as an increase in the ratio of emission at 526 nm versus 476 nm (YFP/ CFP) when CFP is excited by 434 nm light.


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Cell_Biology_Alberts. Incubation of the reporter protein with Abl protein tyrosine kinase in the presence of ATP gave an increase in YFP/CFP emission (Figure Q9–4B). In the absence of ATP or the Abl protein, no FRET occurred. FRET was also eliminated by addition of a tyrosine phosphatase (Figure Q9–4B). Describe as best you can how the reporter protein detects active Abl protein tyrosine kinase. Celis Je, Carter n, simons k et al. (eds) (2005) Cell Biology: a Laboratory handbook, 3rd ed. san Diego: academic press. (Volume 3 of this four-volume set covers the practicalities of most of the current light and electron imaging methods that are used in cell biology.) pawley Bp (ed) (2006) handbook of Biological Confocal Microscopy, 3rd ed. new York: springer science. Wayne r (2014) Light and Video Microscopy. san Diego: academic press.


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Cell_Biology_Alberts. Wayne r (2014) Light and Video Microscopy. san Diego: academic press. Looking at Cells in the Light Microscope adams MC, salmon WC, gupton sL et al. (2003) a high-speed multispectral spinning-disk confocal microscope system for fluorescent speckle microscopy of living cells. Methods 29, 29–41. agard Da, hiraoka Y, shaw p & sedat JW (1989) Fluorescence microscopy in three dimensions. in Methods in Cell Biology, Vol. 30: Fluorescence Microscopy of Living Cells in Culture, part B (DL taylor, Y-L Wang eds). san Diego: academic press. Burnette Dt, sengupta p, Dai Y et al. (2011) Bleaching/blinking assisted localization microscopy for superresolution imaging using standard fluorescent molecules. Proc. Natl Acad. Sci. USA 108, 21081–21086. Chalfie M, tu Y, euskirchen g et al. (1994) green fluorescent protein as a marker for gene expression. Science 263, 802–805.


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Cell_Biology_Alberts. Chalfie M, tu Y, euskirchen g et al. (1994) green fluorescent protein as a marker for gene expression. Science 263, 802–805. giepmans Bn, adams sr, ellisman Mh & tsien rY (2006) the fluorescent toolbox for assessing protein location and function. Science 312, 217–224. harlow e & Lane D (1998) Using antibodies: a Laboratory Manual. Cold spring harbor, nY: Cold spring harbor Laboratory press. hell s (2009) Microscopy and its focal switch. Nat. Methods 6, 24–32. huang B, Babcock h & Zhuang X (2010) Breaking the diffraction barrier: super-resolution imaging of cells. Cell 143, 1047–1058. huang B, Bates M & Zhuang X (2009) super-resolution fluorescence microscopy. Annu. Rev. Biochem. 78, 993–1016. Jaiswai Jk & simon sM (2004) potentials and pitfalls of fluorescent quantum dots for biological imaging. Trends Cell Biol. 14, 497–504.


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Cell_Biology_Alberts. Jaiswai Jk & simon sM (2004) potentials and pitfalls of fluorescent quantum dots for biological imaging. Trends Cell Biol. 14, 497–504. klar ta, Jakobs s, Dyba M et al. (2000) Fluorescence microscopy with diffraction resolution barrier broken by stimulated emission. Proc. Natl Acad. Sci. USA 97, 8206–8210. Lippincott-schwartz J & patterson gh (2003) Development and use of fluorescent protein markers in living cells. Science 300, 87–91. Lippincott-schwartz J, altan-Bonnet n & patterson g (2003) photobleaching and photoactivation: following protein dynamics in living cells. Nat. Cell Biol. 5(suppl), s7–s14. Mcevoy aL, greenfield D, Bates M & Liphardt J (2010) Q&a: single-molecule localization microscopy for biological imaging. BMC Biol. 8, 106. Minsky M (1988) Memoir on inventing the confocal scanning microscope. Scanning 10, 128–138. Miyawaki a, sawano a & kogure t (2003) Lighting up cells: labelling proteins with fluorophores. Nat. Cell Biol. 5(suppl), s1–s7.


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Cell_Biology_Alberts. Miyawaki a, sawano a & kogure t (2003) Lighting up cells: labelling proteins with fluorophores. Nat. Cell Biol. 5(suppl), s1–s7. parton rM & read nD (1999) Calcium and ph imaging in living cells. in Light Microscopy in Biology. a practical approach, 2nd ed. (Lacey aJ ed.). oxford: oxford University press. patterson g, Davidson M, Manley s & Lippincott-schwartz J (2010) superresolution imaging using single-molecule localization. Annu. Rev. Phys. Chem. 61, 345–367. rust MJ, Bates M & Zhuang X (2006) sub-diffraction-limit imaging by stochastic optical reconstruction microscopy (storM). Nat. Methods 3, 793–795. sako Y & Yanagida t (2003) single-molecule visualization in cell biology. Nat. Rev. Mol. Cell Biol. 4(suppl), ss1–ss5. schermelleh L, heintzmann r & Leonhardt h (2010) a guide to super-resolution fluorescence microscopy. J. Cell Biol. 190, 165–175. shaner nC, steinbach pa & tsien rY (2005) a guide to choosing fluorescent proteins. Nat. Methods 2, 905–909.


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Cell_Biology_Alberts. shaner nC, steinbach pa & tsien rY (2005) a guide to choosing fluorescent proteins. Nat. Methods 2, 905–909. sluder g & Wolf De (2007) Digital Microscopy, 3rd ed: Methods in Cell Biology, Vol 81. san Diego: academic press. stephens DJ & allan VJ (2003) Light microscopy techniques for live cell imaging. Science 300, 82–86. tsien rY (2008) Constructing and exploiting the fluorescent protein paintbox. nobel prize Lecture. www.nobelprize.org White Jg, amos WB & Fordham M (1987) an evaluation of confocal versus conventional imaging of biological structures by fluorescence light microscopy. J. Cell Biol. 105, 41–48. Zernike F (1955) how i discovered phase contrast. Science 121, 345–349. Looking at Cells and Molecules in the Electron Microscope allen tD & goldberg MW (1993) high-resolution seM in cell biology. Trends Cell Biol. 3, 205–208. Baumeister W (2002) electron tomography: towards visualizing the molecular organization of the cytoplasm. Curr. Opin. Struct. Biol. 12, 679–684.


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Cell_Biology_Alberts. Baumeister W (2002) electron tomography: towards visualizing the molecular organization of the cytoplasm. Curr. Opin. Struct. Biol. 12, 679–684. Böttcher B, Wynne sa & Crowther ra (1997) Determination of the fold of the core protein of hepatitis B virus by electron cryomicroscopy. Nature 386, 88–91. Dubochet J, adrian M, Chang J-J et al. (1988) Cryo-electron microscopy of vitrified specimens. Q. Rev. Biophys. 21, 129–228. Frank J (2003) electron microscopy of functional ribosome complexes. Biopolymers 68, 223–233. Frank J (2009) single-particle reconstruction of biological macromolecules in electron microscopy—30 years. Quart. Rev. Biophys. 42, 139–158. hayat Ma (2000) principles and techniques of electron Microscopy, 4th ed. Cambridge: Cambridge University press. heuser J (1981) Quick-freeze, deep-etch preparation of samples for 3-D electron microscopy. Trends Biochem. Sci. 6, 64–68.


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Cell_Biology_Alberts. heuser J (1981) Quick-freeze, deep-etch preparation of samples for 3-D electron microscopy. Trends Biochem. Sci. 6, 64–68. Liao M, Cao e, Julius D & Cheng Y (2014) single particle electron cryo-microscopy of a mammalian ion channel. Curr. Opin. Struct. Biol. 27, 1–7. Lucic V, Förster F & Baumeister W (2005) structural studies by electron tomography: from cells to molecules. Annu. Rev. Biochem. 74, 833–865. McDonald kL & auer M (2006) high-pressure freezing, cellular tomography, and structural cell biology. Biotechniques 41, 137–139. Mcintosh Jr (2007) Cellular electron Microscopy. 3rd ed: Methods in Cell Biology, Vol 79. san Diego: academic press. Mcintosh r, nicastro D & Mastronarde D (2005) new views of cells in 3D: an introduction to electron tomography. Trends Cell Biol. 15, 43–51. pease DC & porter kr (1981) electron microscopy and ultramicrotomy. J. Cell Biol. 91, 287s–292s.


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Cell_Biology_Alberts. pease DC & porter kr (1981) electron microscopy and ultramicrotomy. J. Cell Biol. 91, 287s–292s. Unwin pnt & henderson r (1975) Molecular structure determination by electron microscopy of unstained crystalline specimens. J. Mol. Biol. 94, 425–440. Zhou Zh (2008) towards atomic resolution structural determination buy single particle cryo-electron microscopy. Curr. Opin. Struct. Biol. 18, 218–228.


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Cell_Biology_Alberts. Cell membranes are crucial to the life of the cell. The plasma membrane encloses the cell, defines its boundaries, and maintains the essential differences between the cytosol and the extracellular environment. Inside eukaryotic cells, the membranes of the nucleus, endoplasmic reticulum, Golgi apparatus, mitochondria, and other membrane-enclosed organelles maintain the characteristic differences between the contents of each organelle and the cytosol. Ion gradients across membranes, established by the activities of specialized membrane proteins, can be used to synthesize ATP, to drive the transport of selected solutes across the membrane, or, as in nerve and muscle cells, to produce and transmit electrical signals. In all cells, the plasma membrane also contains proteins that act as sensors of external signals, allowing the cell to change its behavior in response to environmental cues, including signals from other cells; these protein sensors, or receptors, transfer information—rather


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Cell_Biology_Alberts. of external signals, allowing the cell to change its behavior in response to environmental cues, including signals from other cells; these protein sensors, or receptors, transfer information—rather than molecules—across the membrane.


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Cell_Biology_Alberts. Despite their differing functions, all biological membranes have a common general structure: each is a very thin film of lipid and protein molecules, held together mainly by noncovalent interactions (Figure 10–1). Cell membranes the lIpID BIlAYer MeMBrAne prOteInS lipid molecule (B) protein molecule lipid bilayer (5 nm) (A) Figure 10–1 Two views of a cell membrane. (A) An electron micrograph of a segment of the plasma membrane of a human red blood cell seen in cross section, showing its bilayer structure. (B) A three-dimensional schematic view of a cell membrane and the general disposition of its lipid and protein constituents. (A, courtesy of Daniel S. Friend.) are dynamic, fluid structures, and most of their molecules move about in the plane of the membrane. The lipid molecules are arranged as a continuous double layer about 5 nm thick. This lipid bilayer provides the basic fluid structure of the membrane and serves as a relatively impermeable barrier to the passage of most


Cell_Biology_Alberts

Cell_Biology_Alberts. are arranged as a continuous double layer about 5 nm thick. This lipid bilayer provides the basic fluid structure of the membrane and serves as a relatively impermeable barrier to the passage of most water-soluble molecules. Most membrane proteins span the lipid bilayer and mediate nearly all of the other functions of the membrane, including the transport of specific molecules across it, and the catalysis of membrane-associated reactions such as ATP synthesis. In the plasma membrane, some transmembrane proteins serve as structural links that connect the cytoskeleton through the lipid bilayer to either the extracellular matrix or an adjacent cell, while others serve as receptors to detect and transduce chemical signals in the cell’s environment. It takes many kinds of membrane proteins to enable a cell to function and interact with its environment, and it is estimated that about 30% of the proteins encoded in an animal’s genome are membrane proteins.


Cell_Biology_Alberts

Cell_Biology_Alberts. In this chapter, we consider the structure and organization of the two main constituents of biological membranes—the lipids and the proteins. Although we focus mainly on the plasma membrane, most concepts discussed apply to the various internal membranes of eukaryotic cells as well. The functions of cell membranes are considered in later chapters: their role in energy conversion and ATP synthesis, for example, is discussed in Chapter 14; their role in the transmembrane transport of small molecules in Chapter 11; and their roles in cell signaling and cell adhesion in Chapters 15 and 19, respectively. In Chapters 12 and 13, we discuss the internal membranes of the cell and the protein traffic through and between them. the lIpID BIlAYer


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Cell_Biology_Alberts. the lIpID BIlAYer The lipid bilayer provides the basic structure for all cell membranes. It is easily seen by electron microscopy, and its bilayer structure is attributable exclusively to the special properties of the lipid molecules, which assemble spontaneously into bilayers even under simple artificial conditions. In this section, we discuss the different types of lipid molecules found in cell membranes and the general properties of lipid bilayers. phosphoglycerides, Sphingolipids, and Sterols Are the Major lipids in cell Membranes


Cell_Biology_Alberts

Cell_Biology_Alberts. phosphoglycerides, Sphingolipids, and Sterols Are the Major lipids in cell Membranes Lipid molecules constitute about 50% of the mass of most animal cell membranes, nearly all of the remainder being protein. There are approximately 5 × 106 lipid molecules in a 1 μm × 1 μm area of lipid bilayer, or about 109 lipid molecules in the plasma membrane of a small animal cell. All of the lipid molecules in cell membranes are amphiphilic—that is, they have a hydrophilic (“water-loving”) or polar end and a hydrophobic (“water-fearing”) or nonpolar end.


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Cell_Biology_Alberts. The most abundant membrane lipids are the phospholipids. These have a polar head group containing a phosphate group and two hydrophobic hydrocarbon tails. In animal, plant, and bacterial cells, the tails are usually fatty acids, and they can differ in length (they normally contain between 14 and 24 carbon atoms). One tail typically has one or more cis-double bonds (that is, it is unsaturated), while the other tail does not (that is, it is saturated). As shown in Figure 10–2, each cis-double bond creates a kink in the tail. Differences in the length and saturation of the fatty acid tails influence how phospholipid molecules pack against one another, thereby affecting the fluidity of the membrane, as we discuss later.


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Cell_Biology_Alberts. The main phospholipids in most animal cell membranes are the phosphoglycerides, which have a three-carbon glycerol backbone (see Figure 10–2). Two long-chain fatty acids are linked through ester bonds to adjacent carbon atoms of the glycerol, and the third carbon atom of the glycerol is attached to a phosphate group, which in turn is linked to one of several types of head group. By combining several different fatty acids and head groups, cells make many different phosphoglycerides. Phosphatidylethanolamine, phosphatidylserine, and phosphatidylcholine are the most abundant ones in mammalian cell membranes (Figure 10–3A–C).


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Cell_Biology_Alberts. Another important class of phospholipids are the sphingolipids, which are built from sphingosine rather than glycerol (Figure10–3D–E). Sphingosine is a long acyl chain with an amino group (NH2) and two hydroxyl groups (OH) at one end. In sphingomyelin, the most common sphingolipid, a fatty acid tail is attached to the amino group, and a phosphocholine group is attached to the terminal hydroxyl group. Together, the phospholipids phosphatidylcholine, phosphatidylethanolamine, phosphatidylserine, and sphingomyelin constitute more than half the mass of lipid in most mammalian cell membranes (see Table 10–1, p. 571). Figure 10–2 The parts of a typical phospholipid molecule. this example is a phosphatidylcholine, represented (A) schematically, (B) by a formula, (c) as a space-filling model (Movie 10.1), and hydrophilicheadhydrophobictails(D) (D) as a symbol.


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Cell_Biology_Alberts. Figure 10–3 Four major phospholipids in mammalian plasma membranes. Different head groups are represented by different colors in the symbols. the lipid molecules shown in (A–c) are phosphoglycerides, which are derived from glycerol. the molecule in (D) is sphingomyelin, which is derived from sphingosine (e) and is therefore a sphingolipid. note that only phosphatidylserine carries a net negative charge, the importance of which we discuss later; the other three are electrically neutral at physiological ph, carrying one positive and one negative charge. Figure 10–4 The structure of cholesterol. polar head group cholesterol is represented (A) by a formula, (B) by a schematic drawing, and (c) as a space-filling model. on their shape, they can do this in either of two ways: they can form spherical micelles, with the tails inward, or they can form double-layered sheets, or bilayers, with the hydrophobic tails sandwiched between the hydrophilic head groups (Figure 10–7).


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Cell_Biology_Alberts. The same forces that drive phospholipids to form bilayers also provide a self-sealing property. A small tear in the bilayer creates a free edge with water; because this is energetically unfavorable, the lipids tend to rearrange spontaneously to eliminate the free edge. (In eukaryotic plasma membranes, the fusion of intracellular vesicles repairs larger tears.) The prohibition of free edges has a profound consequence: the only way for a bilayer to avoid having edges is by closing in on itself and forming a sealed compartment (Figure 10–8). This remarkable


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Cell_Biology_Alberts. In addition to phospholipids, the lipid bilayers in many cell membranes contain glycolipids and cholesterol. Glycolipids resemble sphingolipids, but, instead of a phosphate-linked head group, they have sugars attached. We discuss glycolipids later. Eukaryotic plasma membranes contain especially large amounts of cholesterol—up to one molecule for every phospholipid molecule. Cholesterol is a sterol. It contains a rigid ring structure, to which is attached a single polar hydroxyl group and a short nonpolar hydrocarbon chain (Figure 10–4). The cholesterol molecules orient themselves in the bilayer with their hydroxyl group close to the polar head groups of adjacent phospholipid molecules (Figure 10–5).


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Cell_Biology_Alberts. The shape and amphiphilic nature of the phospholipid molecules cause them to form bilayers spontaneously in aqueous environments. As discussed in Chapter 2, hydrophilic molecules dissolve readily in water because they contain charged groups or uncharged polar groups that can form either favorable electrostatic interactions or hydrogen bonds with water molecules (Figure 10–6A). Hydrophobic molecules, by contrast, are insoluble in water because all, or almost all, of their atoms are uncharged and nonpolar and therefore cannot form energetically favorable interactions with water molecules. If dispersed in water, they force the adjacent water molecules to reorganize into icelike cages that surround the hydrophobic molecule (Figure 10–6B). Because these cage structures are more ordered than the surrounding water, their formation increases the free energy. This free-energy cost is minimized, however, if the hydrophobic molecules (or the hydrophobic portions of amphiphilic molecules) cluster


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Cell_Biology_Alberts. surrounding water, their formation increases the free energy. This free-energy cost is minimized, however, if the hydrophobic molecules (or the hydrophobic portions of amphiphilic molecules) cluster together so that the smallest number of water molecules is affected. 3


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Cell_Biology_Alberts. When amphiphilic molecules are exposed to an aqueous environment, they behave as you would expect from the above discussion. They spontaneously aggregate to bury their hydrophobic tails in the interior, where they are shielded 2 from the water, and they expose their hydrophilic heads to water. Depending Figure 10–5 Cholesterol in a lipid bilayer. Schematic drawing (to scale) of a cholesterol molecule interacting with two phospholipid molecules in one monolayer of a lipid bilayer. behavior, fundamental to the creation of a living cell, follows directly from the shape and amphiphilic nature of the phospholipid molecule. A lipid bilayer also has other characteristics that make it an ideal structure for cell membranes. One of the most important of these is its fluidity, which is crucial to many membrane functions (Movie 10.2). the lipid Bilayer Is a two-dimensional Fluid


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Cell_Biology_Alberts. the lipid Bilayer Is a two-dimensional Fluid Around 1970, researchers first recognized that individual lipid molecules are able to diffuse freely within the plane of a lipid bilayer. The initial demonstration came from studies of synthetic (artificial) lipid bilayers, which can be made in the form of spherical vesicles, called liposomes (Figure 10–9); or in the form of planar bilayers formed across a hole in a partition between two aqueous compartments or on a solid support. Various techniques have been used to measure the motion of individual lipid molecules and their components. One can construct a lipid molecule, for example, with a fluorescent dye or a small gold particle attached to its polar head group and follow the diffusion of even individual molecules in a membrane. Alternatively, one can modify a lipid head group to carry a “spin label,” such as a nitroxide shape of packing of molecule molecules in water


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Cell_Biology_Alberts. Figure 10–7 Packing arrangements of amphiphilic molecules in an aqueous environment. (A) these molecules spontaneously form micelles or bilayers in water, depending on their shape. cone-shaped amphiphilic molecules (above) form micelles, whereas cylinder-shaped amphiphilic molecules such as phospholipids (below) form bilayers. (B) A micelle and a lipid bilayer seen in cross section. note that micelles of amphiphilic molecules are thought to be much more irregular than drawn here (see Figure 10–26c). Figure 10–6 How hydrophilic and hydrophobic molecules interact differently with water. (A) Because acetone is polar, it can form hydrogen bonds (red) and favorable electrostatic interactions (yellow) with water molecules, which are also polar. thus, acetone readily dissolves in water.


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Cell_Biology_Alberts. (B) By contrast, 2-methyl propane is entirely hydrophobic. Because it cannot form favorable interactions with water, it forces adjacent water molecules to reorganize into icelike cage structures, which increases the free energy. this compound is therefore virtually insoluble in water. the symbol δ– indicates a partial negative charge, and δ+ indicates a partial positive charge. polar atoms are shown in color and nonpolar groups are shown in gray. planar phospholipid bilayer with edges exposed to water Figure 10–8 The spontaneous closure of a phospholipid bilayer to form a sealed compartment. the closed structure is stable because it avoids the exposure of the hydrophobic hydrocarbon tails to water, which would be energetically unfavorable.


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Cell_Biology_Alberts. Figure 10–9 Liposomes. (A) An electron micrograph of unfixed, unstained, synthetic phospholipid vesicles—liposomes—in water, which have been rapidly frozen at liquid-nitrogen temperature. (B) A drawing of a small spherical liposome seen in cross section. liposomes are commonly used as model membranes in experimental studies, especially to study incorporated membrane proteins. (A, from p. Frederik and D. hubert, Methods Enzymol. 391:431–448, 2005. With permission from elsevier.) group (=N–O); this contains an unpaired electron whose spin creates a paramagnetic signal that can be detected by electron spin resonance (ESR) spectroscopy, the principles of which are similar to those of nuclear magnetic resonance (NMR), discussed in Chapter 8. The motion and orientation of a spin-labeled lipid in a bilayer can be deduced from the ESR spectrum. Such studies show that phospholipid molecules in synthetic bilayers very rarely migrate from the monolayer (also called a leaflet) on one side to that


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Cell_Biology_Alberts. a bilayer can be deduced from the ESR spectrum. Such studies show that phospholipid molecules in synthetic bilayers very rarely migrate from the monolayer (also called a leaflet) on one side to that on the other. This process, known as “flip-flop,” occurs on a time scale of hours for any individual molecule, although cholesterol is an exception to this rule and can flip-flop rapidly. In contrast, lipid molecules rapidly exchange places with their neighbors within a monolayer (~107 times per second). This gives rise to a rapid lateral diffusion, with a diffusion coefficient (D) of about 10–8 cm2/sec, which means that an average lipid molecule diffuses the length of a large bacterial cell (~2 μm) in about 1 second. These studies have also shown that individual lipid molecules rotate very rapidly about their long axis and have flexible hydrocarbon chains. Computer simulations show that lipid molecules in synthetic bilayers are very disordered, presenting an irregular surface of variously


Cell_Biology_Alberts

Cell_Biology_Alberts. about their long axis and have flexible hydrocarbon chains. Computer simulations show that lipid molecules in synthetic bilayers are very disordered, presenting an irregular surface of variously spaced and oriented head groups to the water phase on either side of the bilayer (Figure 10–10).


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Cell_Biology_Alberts. Similar mobility studies on labeled lipid molecules in isolated biological membranes and in living cells give results similar to those in synthetic bilayers. They demonstrate that the lipid component of a biological membrane is a two-dimensional liquid in which the constituent molecules are free to move laterally. As in synthetic bilayers, individual phospholipid molecules are normally confined to their own monolayer. This confinement creates a problem for their synthesis. Phospholipid molecules are manufactured in only one monolayer of a membrane, mainly in the cytosolic monolayer of the endoplasmic reticulum membrane. If none of these newly made molecules could migrate reasonably promptly to the noncytosolic monolayer, new lipid bilayer could not be made. The problem is solved by a special class of membrane proteins called phospholipid translocators, or flippases, which catalyze the rapid flip-flop of phospholipids from one mono-layer to the other, as discussed in Chapter 12.


Cell_Biology_Alberts

Cell_Biology_Alberts. Despite the fluidity of the lipid bilayer, liposomes do not fuse spontaneously with one another when suspended in water. Fusion does not occur because the polar lipid head groups bind water molecules that need to be displaced for the bilayers of two different liposomes to fuse. The hydration shell that keeps liposomes apart also insulates the many internal membranes in a eukaryotic cell and prevents their uncontrolled fusion, thereby maintaining the compartmental integrity of membrane-enclosed organelles. All cell membrane fusion events


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Cell_Biology_Alberts. Figure 10–10 The mobility of phospholipid molecules in an artificial lipid bilayer. Starting with a model of 100 phosphatidylcholine molecules arranged in a regular bilayer, a computer calculated the position of every atom after 300 picoseconds of simulated time. From these theoretical calculations, a model of the lipid bilayer emerges that accounts for almost all of the measurable properties of a synthetic lipid bilayer, including its thickness, number of lipid molecules per membrane area, depth of water penetration, and unevenness of the two surfaces. note that the tails in one monolayer can interact with those in the other monolayer, if the tails are long enough. (B) the different motions of a lipid molecule in a bilayer. (A, based on S.W. chiu et al., Biophys. J. 69:1230– 1245, 1995. With permission from the Biophysical Society.) are catalyzed by tightly regulated fusion proteins, which force appropriate membranes into tight proximity, squeezing out the water layer that keeps the


Cell_Biology_Alberts

Cell_Biology_Alberts. With permission from the Biophysical Society.) are catalyzed by tightly regulated fusion proteins, which force appropriate membranes into tight proximity, squeezing out the water layer that keeps the bilayers apart, as we discuss in Chapter 13.


Cell_Biology_Alberts

Cell_Biology_Alberts. the Fluidity of a lipid Bilayer Depends on Its composition The fluidity of cell membranes has to be precisely regulated. Certain membrane transport processes and enzyme activities, for example, cease when the bilayer viscosity is experimentally increased beyond a threshold level.


Cell_Biology_Alberts

Cell_Biology_Alberts. The fluidity of a lipid bilayer depends on both its composition and its temperature, as is readily demonstrated in studies of synthetic lipid bilayers. A synthetic bilayer made from a single type of phospholipid changes from a liquid state to a two-dimensional rigid crystalline (or gel) state at a characteristic temperature. This change of state is called a phase transition, and the temperature at which it occurs is lower (that is, the membrane becomes more difficult to freeze) if the hydrocarbon chains are short or have double bonds. A shorter chain length reduces the tendency of the hydrocarbon tails to interact with one another, in both the same and opposite monolayer, and cis-double bonds produce kinks in the chains that make them more difficult to pack together, so that the membrane remains fluid at lower temperatures (Figure 10–11). Bacteria, yeasts, and other organisms whose temperature fluctuates with that of their environment adjust the fatty acid composition of their


Cell_Biology_Alberts

Cell_Biology_Alberts. remains fluid at lower temperatures (Figure 10–11). Bacteria, yeasts, and other organisms whose temperature fluctuates with that of their environment adjust the fatty acid composition of their membrane lipids to maintain a relatively constant fluidity. As the temperature falls, for instance, the cells of those organisms synthesize fatty acids with more cis-double bonds, thereby avoiding the decrease in bilayer fluidity that would otherwise result from the temperature drop.


Cell_Biology_Alberts

Cell_Biology_Alberts. Cholesterol modulates the properties of lipid bilayers. When mixed with phospholipids, it enhances the permeability-barrier properties of the lipid bilayer. Cholesterol inserts into the bilayer with its hydroxyl group close to the polar head groups of the phospholipids, so that its rigid, platelike steroid rings interact with— and partly immobilize—those regions of the hydrocarbon chains closest to the polar head groups (see Figure 10–5 and Movie 10.3). By decreasing the mobility of the first few CH2 groups of the chains of the phospholipid molecules, cholesterol makes the lipid bilayer less deformable in this region and thereby decreases the permeability of the bilayer to small water-soluble molecules. Although cholesterol tightens the packing of the lipids in a bilayer, it does not make membranes any less fluid. At the high concentrations found in most eukaryotic plasma membranes, cholesterol also prevents the hydrocarbon chains from coming together and crystallizing.


Cell_Biology_Alberts

Cell_Biology_Alberts. Table 10–1 compares the lipid compositions of several biological membranes. Note that bacterial plasma membranes are often composed of one main type of phospholipid and contain no cholesterol. In archaea, lipids usually contain unsaturated saturated hydrocarbon chains hydrocarbon chains with cis-double bonds Figure 10–11 The influence of cisdouble bonds in hydrocarbon chains. the double bonds make it more difficult to pack the chains together, thereby making the lipid bilayer more difficult to freeze. In addition, because the hydrocarbon chains of unsaturated lipids are more spread apart, lipid bilayers containing them are thinner than bilayers formed exclusively from saturated lipids.


Cell_Biology_Alberts

Cell_Biology_Alberts. 20–25-carbon-long prenyl chains instead of fatty acids; prenyl and fatty acid chains are similarly hydrophobic and flexible (see Figure 10–20F); in thermophilic archaea, the longest lipid chains span both leaflets, making the membrane particularly stable to heat. Thus, lipid bilayers can be built from molecules with similar features but different molecular designs. The plasma membranes of most eukaryotic cells are more varied than those of prokaryotes and archaea, not only in containing large amounts of cholesterol but also in containing a mixture of different phospholipids.


Cell_Biology_Alberts

Cell_Biology_Alberts. Analysis of membrane lipids by mass spectrometry has revealed that the lipid composition of a typical eukaryotic cell membrane is much more complex than originally thought. These membranes contain a bewildering variety of perhaps 500–2000 different lipid species with even the simple plasma membrane of a red blood cell containing well over 150. While some of this complexity reflects the combinatorial variation in head groups, hydrocarbon chain lengths, and desaturation of the major phospholipid classes, some membranes also contain many structurally distinct minor lipids, at least some of which have important functions. The inositol phospholipids, for example, are present in small quantities in animal cell membranes and have crucial functions in guiding membrane traffic and in cell signaling (discussed in Chapters 13 and 15, respectively). Their local synthesis and destruction are regulated by a large number of enzymes, which create both small intracellular signaling molecules and lipid


Cell_Biology_Alberts

Cell_Biology_Alberts. (discussed in Chapters 13 and 15, respectively). Their local synthesis and destruction are regulated by a large number of enzymes, which create both small intracellular signaling molecules and lipid docking sites on membranes that recruit specific proteins from the cytosol, as we discuss later.


Cell_Biology_Alberts

Cell_Biology_Alberts. Despite their Fluidity, lipid Bilayers can Form Domains of Different compositions Because a lipid bilayer is a two-dimensional fluid, we might expect most types of lipid molecules in it to be well mixed and randomly distributed in their own monolayer. The van der Waals attractive forces between neighboring hydrocarbon tails are not selective enough to hold groups of phospholipid molecules together. With certain lipid mixtures in artificial bilayers, however, one can observe phase segregations in which specific lipids come together in separate domains (Figure 10–12).


Cell_Biology_Alberts

Cell_Biology_Alberts. There has been a long debate among cell biologists about whether the lipid molecules in the plasma membrane of living cells similarly segregate into specialized domains, called lipid rafts. Although many lipids and membrane proteins are not distributed uniformly, large-scale lipid phase segregations are rarely seen in living cell membranes. Instead, specific membrane proteins and lipids are seen to concentrate in a more temporary, dynamic fashion facilitated by protein– protein interactions that allow the transient formation of specialized membrane regions (Figure 10–13). Such clusters can be tiny nanoclusters on a scale of a few molecules, or larger assemblies that can be seen with electron microscopy, such as the caveolae involved in endocytosis (discussed in Chapter 13). The tendency of mixtures of lipids to undergo phase partitioning, as seen in artificial bilayers (see Figure 10–12), may help create rafts in living cell membranes—organizing and concentrating membrane proteins


Cell_Biology_Alberts

Cell_Biology_Alberts. of mixtures of lipids to undergo phase partitioning, as seen in artificial bilayers (see Figure 10–12), may help create rafts in living cell membranes—organizing and concentrating membrane proteins either for transport in membrane vesicles


Cell_Biology_Alberts

Cell_Biology_Alberts. Figure 10–12 Lateral phase separation in artificial lipid bilayers. (A) giant liposomes produced from a 1:1 mixture of phosphatidylcholine and sphingomyelin form uniform bilayers. (B) By contrast, liposomes produced from a 1:1:1 mixture of phosphatidylcholine, sphingomyelin, and cholesterol form bilayers with two separate phases. the liposomes are stained with trace concentrations of a fluorescent dye that preferentially partitions into one of the two phases. the average size of the domains formed in these giant artificial liposomes is much larger than that expected in cell membranes, where “lipid rafts” (see text) may be as small as a few nanometers in diameter. (A, from n. Kahya et al., J. Struct. Biol. 147:77–89, 2004. With permission from elsevier; B, courtesy of petra Schwille.) (discussed in Chapter 13) or for working together in protein assemblies, such as when they convert extracellular signals into intracellular ones (discussed in Chapter 15).


Cell_Biology_Alberts

Cell_Biology_Alberts. Most cells store an excess of lipids in lipid droplets, from where they can be retrieved as building blocks for membrane synthesis or as a food source. Fat cells, or adipocytes, are specialized for lipid storage. They contain a giant lipid droplet that fills up most of their cytoplasm. Most other cells have many smaller lipid droplets, the number and size varying with the cell’s metabolic state. Fatty acids can be liberated from lipid droplets on demand and exported to other cells through the bloodstream. Lipid droplets store neutral lipids, such as triacylglycerols and cholesterol esters, which are synthesized from fatty acids and cholesterol by enzymes in the endoplasmic reticulum membrane. Because these lipids do not contain hydrophilic head groups, they are exclusively hydrophobic molecules, and therefore aggregate into three-dimensional droplets rather than into bilayers.


Cell_Biology_Alberts

Cell_Biology_Alberts. Lipid droplets are unique organelles in that they are surrounded by a single monolayer of phospholipids, which contains a large variety of proteins. Some of the proteins are enzymes involved in lipid metabolism, but the functions of most are unknown. Lipid droplets form rapidly when cells are exposed to high concentrations of fatty acids. They are thought to form from discrete regions of the endoplasmic reticulum membrane where many enzymes of lipid metabolism are concentrated. Figure 10–14 shows one model of how lipid droplets may form and acquire their surrounding monolayer of phospholipids and proteins. the Asymmetry of the lipid Bilayer Is Functionally Important The lipid compositions of the two monolayers of the lipid bilayer in many membranes are strikingly different. In the human red blood cell (erythrocyte) membrane, for example, almost all of the phospholipid molecules that have cho-line—(CH3)3N+CH2CH2OH—in their head group (phosphatidylcholine and


Cell_Biology_Alberts

Cell_Biology_Alberts. Figure 10–13 a model of a raft domain. Weak protein–protein, protein–lipid, and lipid–lipid interactions reinforce one another to partition the interacting components into raft domains. cholesterol, sphingolipids, glycolipids, glycosylphosphatidylinositol (gpI)-anchored proteins, and some transmembrane proteins are enriched in these domains. note that because of their composition, raft domains have an increased membrane thickness.We discuss glycolipids, gpI-anchored proteins, and oligosaccharide linkers later. (Adapted from D. lingwood and K. Simons, Science 327:46–50, 2010.)


Cell_Biology_Alberts

Cell_Biology_Alberts. Figure 10–14 a model for the formation of lipid droplets. neutral lipids are deposited between the two monolayers of the endoplasmic reticulum membrane. there, they aggregate into a three-dimensional droplet, which buds and pinches off from the endoplasmic reticulum membrane as a unique organelle, surrounded by a single monolayer of phospholipids and associated proteins. (Adapted from S. Martin and r.g. parton, Nat. Rev. Mol. Cell Biol. 7:373–378, 2006. With permission from Macmillan publishers ltd.) sphingomyelin) are in the outer monolayer, whereas almost all that contain a terminal primary amino group (phosphatidylethanolamine and phosphatidylserine) are in the inner monolayer (Figure 10–15). Because the negatively charged phosphatidylserine is located in the inner monolayer, there is a significant difference in charge between the two halves of the bilayer. We discuss in Chapter 12 how membrane-bound phospholipid translocators generate and maintain lipid asymmetry.


Cell_Biology_Alberts

Cell_Biology_Alberts. Lipid asymmetry is functionally important, especially in converting extracellular signals into intracellular ones (discussed in Chapter 15). Many cytosolic proteins bind to specific lipid head groups found in the cytosolic monolayer of the lipid bilayer. The enzyme protein kinase C (PKC), for example, which is activated in response to various extracellular signals, binds to the cytosolic face of the plasma membrane, where phosphatidylserine is concentrated, and requires this negatively charged phospholipid for its activity.


Cell_Biology_Alberts

Cell_Biology_Alberts. In other cases, specific lipid head groups must first be modified to create pro-tein-binding sites at a particular time and place. One example is phosphatidylinositol (PI), one of the minor phospholipids that are concentrated in the cytosolic monolayer of cell membranes (see Figure 13–10A–C). Various lipid kinases can add phosphate groups at distinct positions on the inositol ring, creating binding sites that recruit specific proteins from the cytosol to the membrane. An important example of such a lipid kinase is phosphoinositide 3-kinase (PI 3-kinase), which is activated in response to extracellular signals and helps to recruit specific intracellular signaling proteins to the cytosolic face of the plasma membrane (see Figure 15–53). Similar lipid kinases phosphorylate inositol phospholipids in intracellular membranes and thereby help to recruit proteins that guide membrane transport.


Cell_Biology_Alberts

Cell_Biology_Alberts. Phospholipids in the plasma membrane are used in yet another way to convert extracellular signals into intracellular ones. The plasma membrane contains various phospholipases that are activated by extracellular signals to cleave specific phospholipid molecules, generating fragments of these molecules that act as short-lived intracellular mediators. Phospholipase C, for example, cleaves an inositol phospholipid in the cytosolic monolayer of the plasma membrane to generate two fragments, one of which remains in the membrane and helps activate protein kinase C, while the other is released into the cytosol and stimulates the release of Ca2+ from the endoplasmic reticulum (see Figure 15–28).


Cell_Biology_Alberts

Cell_Biology_Alberts. Animals exploit the phospholipid asymmetry of their plasma membranes to distinguish between live and dead cells. When animal cells undergo apoptosis (a form of programmed cell death, discussed in Chapter 18), phosphatidylserine, which is normally confined to the cytosolic (or inner) monolayer of the plasma membrane lipid bilayer, rapidly translocates to the extracellular (or outer) mono-layer. The phosphatidylserine exposed on the cell surface signals neighboring cells, such as macrophages, to phagocytose the dead cell and digest it. The trans-location of the phosphatidylserine in apoptotic cells is thought to occur by two mechanisms: 1. The phospholipid translocator that normally transports this lipid from the outer monolayer to the inner monolayer is inactivated. 2. A “scramblase” that transfers phospholipids nonspecifically in both directions between the two monolayers is activated.


Cell_Biology_Alberts

Cell_Biology_Alberts. 2. A “scramblase” that transfers phospholipids nonspecifically in both directions between the two monolayers is activated. Figure 10–15 The asymmetrical distribution of phospholipids and glycolipids in the lipid bilayer of human red blood cells. the colors used for the phospholipid head groups are those introduced in Figure 10–3. In addition, glycolipids are drawn with hexagonal polar head groups (blue). cholesterol (not shown) is distributed roughly equally in both monolayers. glycolipids Are Found on the Surface of All eukaryotic plasma Membranes


Cell_Biology_Alberts

Cell_Biology_Alberts. Sugar-containing lipid molecules called glycolipids have the most extreme asymmetry in their membrane distribution: these molecules, whether in the plasma membrane or in intracellular membranes, are found exclusively in the monolayer facing away from the cytosol. In animal cells, they are made from sphingosine, just like sphingomyelin (see Figure 10–3). These intriguing molecules tend to self-associate, partly through hydrogen bonds between their sugars and partly through van der Waals forces between their long and straight hydrocarbon chains, which causes them to partition preferentially into lipid raft phases (see Figure 10–13). The asymmetric distribution of glycolipids in the bilayer results from the addition of sugar groups to the lipid molecules in the lumen of the Golgi apparatus. Thus, the compartment in which they are manufactured is topologically equivalent to the exterior of the cell (discussed in Chapter 12). As they are delivered to the plasma membrane, the sugar groups


Cell_Biology_Alberts

Cell_Biology_Alberts. Thus, the compartment in which they are manufactured is topologically equivalent to the exterior of the cell (discussed in Chapter 12). As they are delivered to the plasma membrane, the sugar groups are exposed at the cell surface (see Figure 10–15), where they have important roles in interactions of the cell with its surroundings.


Cell_Biology_Alberts

Cell_Biology_Alberts. Glycolipids probably occur in all eukaryotic cell plasma membranes, where they generally constitute about 5% of the lipid molecules in the outer monolayer. They are also found in some intracellular membranes. The most complex of the glycolipids, the gangliosides, contain oligosaccharides with one or more sialic acid moieties, which give gangliosides a net negative charge (Figure 10–16). The most abundant of the more than 40 different gangliosides that have been identified are in the plasma membrane of nerve cells, where gangliosides constitute 5–10% of the total lipid mass; they are also found in much smaller quantities in other cell types.


Cell_Biology_Alberts

Cell_Biology_Alberts. Hints as to the functions of glycolipids come from their localization. In the plasma membrane of epithelial cells, for example, glycolipids are confined to the exposed apical surface, where they may help to protect the membrane against the harsh conditions frequently found there (such as low pH and high concentrations of degradative enzymes). Charged glycolipids, such as gangliosides, may be important because of their electrical effects: their presence alters the electrical field across the membrane and the concentrations of ions—especially Ca2+—at the membrane surface. Glycolipids also function in cell-recognition processes, Figure 10–16 Glycolipid molecules.


Cell_Biology_Alberts

Cell_Biology_Alberts. Figure 10–16 Glycolipid molecules. (A) galactocerebroside is called a neutral glycolipid because the sugar that forms its head group is uncharged. (B) A ganglioside always contains one or more negatively charged sialic acid moiety. there are various types of sialic acid; in human cells, it is mostly N-acetylneuraminic acid, or nAnA), whose structure is shown in (c). Whereas in bacteria and plants almost all glycolipids are derived from glycerol, as are most phospholipids, in animal cells almost all glycolipids are based on sphingosine, as is the case for sphingomyelin (see Figure 10–3). gal = galactose; glc = glucose, galnAc = N-acetylgalactosamine; these three sugars are uncharged.


Cell_Biology_Alberts

Cell_Biology_Alberts. in which membrane-bound carbohydrate-binding proteins (lectins) bind to the sugar groups on both glycolipids and glycoproteins in the process of cell–cell adhesion (discussed in Chapter 19). Mutant mice that are deficient in all of their complex gangliosides show abnormalities in the nervous system, including axonal degeneration and reduced myelination.


Cell_Biology_Alberts

Cell_Biology_Alberts. Some glycolipids provide entry points for certain bacterial toxins and viruses. The ganglioside GM1 (see Figure 10–16), for example, acts as a cell-surface receptor for the bacterial toxin that causes the debilitating diarrhea of cholera. Cholera toxin binds to and enters only those cells that have GM1 on their surface, including intestinal epithelial cells. Its entry into a cell leads to a prolonged increase in the concentration of intracellular cyclic AMP (discussed in Chapter 15), which in turn causes a large efflux of Cl–, leading to the secretion of Na+, K+, HCO3–, and water into the intestine. Polyomaviruses also enter the cell after binding initially to gangliosides.


Cell_Biology_Alberts

Cell_Biology_Alberts. Biological membranes consist of a continuous double layer of lipid molecules in which membrane proteins are embedded. This lipid bilayer is fluid, with individual lipid molecules able to diffuse rapidly within their own monolayer. The membrane lipid molecules are amphiphilic. When placed in water, they assemble spontaneously into bilayers, which form sealed compartments.


Cell_Biology_Alberts

Cell_Biology_Alberts. Although cell membranes can contain hundreds of different lipid species, the plasma membrane in animal cells contains three major classes—phospholipids, cholesterol, and glycolipids. Because of their different backbone structure, phospholipids fall into two subclasses—phosphoglycerides and sphingolipids. The lipid compositions of the inner and outer monolayers are different, reflecting the different functions of the two faces of a cell membrane. Different mixtures of lipids are found in the membranes of cells of different types, as well as in the various membranes of a single eukaryotic cell. Inositol phospholipids are a minor class of phospholipids, which in the cytosolic leaflet of the plasma membrane lipid bilayer play an important part in cell signaling: in response to extracellular signals, specific lipid kinases phosphorylate the head groups of these lipids to form docking sites for cytosolic signaling proteins, whereas specific phospholipases cleave certain inositol


Cell_Biology_Alberts

Cell_Biology_Alberts. signals, specific lipid kinases phosphorylate the head groups of these lipids to form docking sites for cytosolic signaling proteins, whereas specific phospholipases cleave certain inositol phospholipids to generate small intracellular signaling molecules.


Cell_Biology_Alberts

Cell_Biology_Alberts. Although the lipid bilayer provides the basic structure of biological membranes, the membrane proteins perform most of the membrane’s specific tasks and therefore give each type of cell membrane its characteristic functional properties. Accordingly, the amounts and types of proteins in a membrane are highly variable. In the myelin membrane, which serves mainly as electrical insulation for nerve-cell axons, less than 25% of the membrane mass is protein. By contrast, in the membranes involved in ATP production (such as the internal membranes of mitochondria and chloroplasts), approximately 75% is protein. A typical plasma membrane is somewhere in between, with protein accounting for about half of its mass. Because lipid molecules are small compared with protein molecules, however, there are always many more lipid molecules than protein molecules in cell membranes—about 50 lipid molecules for each protein molecule in cell membranes that are 50% protein by mass. Membrane proteins vary


Cell_Biology_Alberts

Cell_Biology_Alberts. are always many more lipid molecules than protein molecules in cell membranes—about 50 lipid molecules for each protein molecule in cell membranes that are 50% protein by mass. Membrane proteins vary widely in structure and in the way they associate with the lipid bilayer, which reflects their diverse functions.


Cell_Biology_Alberts

Cell_Biology_Alberts. Membrane proteins can Be Associated with the lipid Bilayer in Various Ways Figure 10–17 shows the different ways in which proteins can associate with the membrane. Like their lipid neighbors, membrane proteins are amphiphilic, having hydrophobic and hydrophilic regions. Many membrane proteins extend through the lipid bilayer, and hence are called transmembrane proteins, with part of their mass on either side (Figure 10–17, examples 1, 2, and 3). Their hydrophobic regions pass through the membrane and interact with the hydrophobic tails of the lipid molecules in the interior of the bilayer, where they are sequestered away from water. Their hydrophilic regions are exposed to water on either side of the membrane. The covalent attachment of a fatty acid chain that inserts into the cytosolic monolayer of the lipid bilayer increases the hydrophobicity of some of these transmembrane proteins (see Figure 10–17, example 1).


Cell_Biology_Alberts

Cell_Biology_Alberts. Other membrane proteins are located entirely in the cytosol and are attached to the cytosolic monolayer of the lipid bilayer, either by an amphiphilic α helix exposed on the surface of the protein (Figure 10–17, example 4) or by one or more covalently attached lipid chains (Figure 10–17, example 5). Yet other membrane proteins are entirely exposed at the external cell surface, being attached to the lipid bilayer only by a covalent linkage (via a specific oligosaccharide) to a lipid anchor in the outer monolayer of the plasma membrane (Figure 10–17, example 6).