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Cell_Biology_Alberts. Microtubule dynamics, like those of actin filaments, are profoundly influenced by the binding and hydrolysis of nucleotide—GTP in this case. GTP hydrolysis occurs only within the β-tubulin subunit of the tubulin dimer. It proceeds very slowly in free tubulin subunits but is accelerated when they are incorporated into microtubules. Following GTP hydrolysis, the free phosphate group is released and the GDP remains bound to β-tubulin within the microtubule lattice. Thus, as in the case of actin filaments, two different types of microtubule structures can exist, one with the “T form” of the nucleotide bound (GTP) and one with the “D form” bound (GDP). The energy of nucleotide hydrolysis is stored as elastic strain in the polymer lattice, making the free-energy change for dissociation of a subunit from the D-form polymer more negative than the free-energy change for dissociation of a subunit from the T-form polymer. In consequence, the ratio of koff/kon for GDP-tubulin (its critical


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Cell_Biology_Alberts. a subunit from the D-form polymer more negative than the free-energy change for dissociation of a subunit from the T-form polymer. In consequence, the ratio of koff/kon for GDP-tubulin (its critical concentration [Cc(D)]) is much higher than that of GTP-tubulin. Thus, under physiological conditions, GTP-tubulin tends to polymerize and GDP-tubulin to depolymerize.


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Cell_Biology_Alberts. Whether the tubulin subunits at the very end of a microtubule are in the T or the D form depends on the relative rates of GTP hydrolysis and tubulin addition. If the rate of subunit addition is high—and thus the filament is growing rapidly— then it is likely that a new subunit will be added to the polymer before the nucleotide in the previously added subunit has been hydrolyzed. In this case, the tip of the polymer remains in the T form, forming a GTP cap. However, if the rate of subunit addition is low, hydrolysis may occur before the next subunit is added, and the tip of the filament will then be in the D form. If GTP-tubulin subunits assemble at the end of the microtubule at a rate similar to the rate of GTP hydrolysis, then hydrolysis will sometimes “catch up” with the rate of subunit addition and transform the end to a D form. This transformation is sudden and random, with a certain probability per unit time that depends on the concentration of free GTP-tubulin subunits.


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Cell_Biology_Alberts. Suppose that the concentration of free tubulin is intermediate between the critical concentration for a T-form end and the critical concentration for a D-form end (that is, above the concentration necessary for T-form assembly, but below that for the D form). Now, any end that happens to be in the T form will grow, whereas any end that happens to be in the D form will shrink. On a single microtubule, an end might grow for a certain length of time in a T form, but then suddenly change to the D form and begin to shrink rapidly, even while the free subunit concentration is held constant. At some later time, it might then regain a T-form end and begin to grow again. This rapid interconversion between a growing and shrinking state, at a uniform free subunit concentration, is called dynamic instability (Figure 16–44A and Figure 16–45; see Panel 16–2). The change from growth to shrinkage is called a catastrophe, while the change to growth is called a rescue.


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Cell_Biology_Alberts. In a population of microtubules, at any instant some of the ends are in the T form and some are in the D form, with the ratio depending on the hydrolysis rate and the free subunit concentration. In vitro, the structural difference between a T-form end and a D-form end is dramatic. Tubulin subunits with GTP bound to rapid growth with GTP-capped end random loss of GTP cap rapid shrinkage regain of GTP cap rapid growth with GTP-capped end etc. 50 nmGTP-tubulin dimer ˜°exchangeable GTP straight protoflament GTP HYDROLYSIS CHANGES SUBUNIT CONFORMATION AND WEAKENS BOND IN THE POLYMER DEPOLYMERIZATION GDP-GTP EXCHANGE (A) GROWING SHRINKING CATASTROPHE RESCUE GTP cap less stable region of microtubule containing GDP-tubulin dimers curved protoflament GDP-tubulin dimer GTPGTPGDPGDPGDPGDPGDPGDPGTPGTPGTP


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Cell_Biology_Alberts. Figure 16–44 Dynamic instability due to the structural differences between a growing and a shrinking microtubule end. (A) If the free tubulin concentration in solution is between the critical concentrations of the GTPand GDP-bound forms, a single microtubule end may undergo transitions between a growing state and a shrinking state. A growing microtubule has GTP-containing subunits at its end, forming a GTP cap. If nucleotide hydrolysis proceeds more rapidly than subunit addition, the cap is lost and the microtubule begins to shrink, an event called a “catastrophe.” But GTP-containing subunits may still add to the shrinking end, and if enough add to form a new cap, then microtubule growth resumes, an event called “rescue.” (B) Model for the structural consequences of GTP hydrolysis in the microtubule lattice. The addition of GTP-containing tubulin subunits to the end of a protofilament causes the end to grow in a linear conformation that can readily pack into the cylindrical wall of


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Cell_Biology_Alberts. microtubule lattice. The addition of GTP-containing tubulin subunits to the end of a protofilament causes the end to grow in a linear conformation that can readily pack into the cylindrical wall of the microtubule. Hydrolysis of GTP after assembly changes the conformation of the subunits and tends to force the protofilament into a curved shape that is less able to pack into the microtubule wall. (C) In an intact microtubule, protofilaments made from GDP-containing subunits are forced into a linear conformation by the many lateral bonds within the microtubule wall, given a stable cap of GTP-containing subunits. Loss of the GTP cap, however, allows the GDP-containing protofilaments to relax into their more curved conformation. This leads to a progressive disruption of the microtubule. Above the drawings of a growing and a shrinking microtubule, electron micrographs show actual microtubules in each of these two states. Note particularly the curling, disintegrating GDP-containing


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Cell_Biology_Alberts. Above the drawings of a growing and a shrinking microtubule, electron micrographs show actual microtubules in each of these two states. Note particularly the curling, disintegrating GDP-containing protofilaments at the end of the shrinking microtubule. (C, from E.M. Mandelkow,


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Cell_Biology_Alberts. E. Mandelkow and R.A. Milligan, J. Cell Biol. 114:977–991, 1991. With permission from The Rockefeller University Press.) the β-monomer produce straight protofilaments that make strong and regular lateral contacts with one another. But the hydrolysis of GTP to GDP is associated with a subtle conformational change in the protein, which makes the protofilaments curved (Figure 16–44B). On a rapidly growing microtubule, the GTP cap is thought to constrain the curvature of the protofilaments, and the ends appear straight. But when the terminal subunits have hydrolyzed their nucleotides, this constraint is removed, and the curved protofilaments spring apart. This cooperative release of the energy of hydrolysis stored in the microtubule lattice causes the curled protofilaments to peel off rapidly, and curved oligomers of GDP-containing tubulin are seen near the ends of depolymerizing microtubules (Figure 16–44C).


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Cell_Biology_Alberts. Chemical compounds that impair polymerization or depolymerization of micro-tubules are powerful tools for investigating the roles of these polymers in cells. Whereas colchicine and nocodazole interact with tubulin subunits and lead to microtubule depolymerization, Taxol binds to and stabilizes microtubules, causing a net increase in tubulin polymerization (see Table 16–1). Drugs like these have a rapid and profound effect on the organization of the microtubules in living cells. Both microtubule-depolymerizing drugs (such as nocodazole) and microtubule-polymerizing drugs (such as Taxol) preferentially kill dividing cells, since microtubule dynamics are crucial for correct function of the mitotic spindle (discussed in Chapter 17). Some of these drugs efficiently kill certain types of tumor cells in a human patient, although not without toxicity to rapidly dividing normal cells, including those in the bone marrow, intestine, and hair follicles. Taxol in particular has been widely used to


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Cell_Biology_Alberts. in a human patient, although not without toxicity to rapidly dividing normal cells, including those in the bone marrow, intestine, and hair follicles. Taxol in particular has been widely used to treat cancers of the breast and lung, and it is frequently successful in treatment of tumors that are resistant to other chemotherapeutic agents.


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Cell_Biology_Alberts. Because formation of a microtubule requires the interaction of many tubulin heterodimers, the concentration of tubulin subunits required for spontaneous nucleation of microtubules is very high. Microtubule nucleation therefore requires help from other factors. While αand β-tubulins are the regular building blocks of microtubules, another type of tubulin, called γ-tubulin, is present in much smaller amounts than αand β-tubulin and is involved in the nucleation of microtubule growth in organisms ranging from yeasts to humans. Microtubules are generally nucleated from a specific intracellular location known as a microtubule-organizing center (MTOC) where γ-tubulin is most enriched. Nucleation in many cases depends on the γ-tubulin ring complex (γ-TuRC). Within this complex, two accessory proteins bind directly to the γ-tubulin, along with several other proteins that help create a spiral ring of γ-tubulin molecules, which serves as a template that creates a microtubule with 13


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Cell_Biology_Alberts. proteins bind directly to the γ-tubulin, along with several other proteins that help create a spiral ring of γ-tubulin molecules, which serves as a template that creates a microtubule with 13 protofilaments (Figure 16–46).


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Cell_Biology_Alberts. Figure 16–45 Direct observation of the dynamic instability of microtubules in a living cell. Microtubules in a newt lung epithelial cell were observed after the cell was injected with a small amount of rhodamine-labeled tubulin. Notice the dynamic instability of microtubules at the edge of the cell. Four individual microtubules are highlighted for clarity; each of these shows alternating shrinkage and growth (Movie 16.7). (Courtesy of Wendy C. Salmon and Clare Waterman-Storer.) schematic of ˛-tubulin seven copies of


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Cell_Biology_Alberts. Figure 16–46 Microtubule nucleation by the γ-tubulin ring complex. (A) Two copies of γ-tubulin associate with a pair of accessory proteins to form the γ-tubulin small complex (γ-TuSC). This image was generated by high-resolution electron microscopy of individual purified complexes. (B) Seven copies of the γ-TuSC associate to form a spiral structure in which the last γ-tubulin lies beneath the first, resulting in 13 exposed γ-tubulin subunits in a circular orientation that matches the orientation of the 13 protofilaments in a microtubule. (C) In many cell types, the γ-TuSC spiral associates with additional accessory proteins to form the γ-tubulin ring complex (γ-TuRC), which is likely to nucleate the minus end of a microtubule as shown here. Note the longitudinal discontinuity between two protofilaments, which results from the spiral orientation of the γ-tubulin subunits. Microtubules often have one such “seam” breaking the otherwise uniform helical packing of the protofilaments. (A


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Cell_Biology_Alberts. protofilaments, which results from the spiral orientation of the γ-tubulin subunits. Microtubules often have one such “seam” breaking the otherwise uniform helical packing of the protofilaments. (A and B, from


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Cell_Biology_Alberts. J.M. Kollman et al., Nature 466:879–883, 2010. With permission from Macmillan Publishers Ltd.) Microtubules Emanate from the Centrosome in Animal Cells Many animal cells have a single, well-defined MTOC called the centrosome, which is located near the nucleus and from which microtubules are nucleated at their minus ends, so the plus ends point outward and continuously grow and shrink, probing the entire three-dimensional volume of the cell. A centrosome typically recruits more than fifty copies of γ-TuRC. In addition, γ-TuRC molecules are found in the cytoplasm, and centrosomes are not absolutely required for microtubule nucleation, since destroying them with a laser pulse does not prevent microtubule nucleation elsewhere in the cell. A variety of proteins have been identified that anchor γ-TuRC to the centrosome, but mechanisms that activate microtubule nucleation at MTOCs and at other sites in the cell are poorly understood.


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Cell_Biology_Alberts. Embedded in the centrosome are the centrioles, a pair of cylindrical structures arranged at right angles to each other in an L-shaped configuration (Figure 16–47). A centriole consists of a cylindrical array of short, modified microtubules arranged into a barrel shape with striking ninefold symmetry (Figure 16–48). Together with a large number of accessory proteins, the centrioles organize the pericentriolar material, where microtubule nucleation takes place. As described in Chapter 17, the centrosome duplicates and splits into two parts before mitosis, each containing a duplicated centriole pair. The two centrosomes move to opposite sides of the nucleus when mitosis begins, and they form the two poles of the mitotic spindle (see Panel 17–1).


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Cell_Biology_Alberts. Microtubule organization varies widely among different species and cell types. In budding yeast, microtubules are nucleated at an MTOC that is embedded in the nuclear envelope as a small, multilayered structure called the spindle pole body, also found in other fungi and diatoms. Higher-plant cells appear to nucleate microtubules at sites distributed all around the nuclear envelope and at the cell cortex. Neither fungi nor most plant cells contain centrioles. Despite these differences, all these cells seem to use γ-tubulin to nucleate their microtubules. In cultured animal cells, the aster-like configuration of microtubules is robust, with dynamic plus ends pointing outward toward the cell periphery and stable minus ends collected near the nucleus. The system of microtubules radiating from pair of ++ + (A) (B) of the centrosome


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Cell_Biology_Alberts. Figure 16–47 The centrosome. (A) The centrosome is the major MTOC of animal cells. Located in the cytoplasm next to the nucleus, it consists of an amorphous matrix of fibrous proteins to which the γ-tubulin ring complexes that nucleate microtubule growth are attached. This matrix is organized by a pair of centrioles, as described in the text. (B) A centrosome with attached microtubules. The minus end of each microtubule is embedded in the centrosome, having grown from a γ-tubulin ring complex, whereas the plus end of each microtubule is free in the cytoplasm. (C) In a reconstructed image of the MTOC from a C. elegans cell, a dense thicket of microtubules can be seen emanating from the centrosome. (C, from E.T. O’Toole et al., J. Cell Biol.


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Cell_Biology_Alberts. 163:451–456, 2003. With permission from the authors.) the centrosome acts as a device to survey the outlying regions of the cell and to position the centrosome at its center. Even in an isolated cell fragment lacking the centrosome, dynamic microtubules arrange themselves into a star-shaped array with the microtubule minus ends clustered at the center by minus-end-binding proteins (Figure 16–49). This ability of the microtubule cytoskeleton to find the center of the cell establishes a general coordinate system, which is then used to position many organelles within the cell. Highly differentiated cells with complex morphologies such as neurons, muscles, and epithelial cells must use additional measuring mechanisms to establish their more elaborate internal coordinate systems. Thus, for example, when an epithelial cell forms cell–cell junctions and becomes highly polarized, the microtubule minus ends move to a region near the


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Cell_Biology_Alberts. Figure 16–48 A pair of centrioles in the centrosome. (A) An electron micrograph of a thin section of an isolated centrosome showing the mother centriole with its distal appendages and the adjacent daughter centriole, which formed through a duplication event during S phase (see Figure 17–26). In the centrosome, the centriole pair is surrounded by a dense matrix of pericentriolar material from which microtubules nucleate. Centrioles also function as basal bodies to nucleate the formation of ciliary axonemes (see Figure 16–68). (B) Electron micrograph of a cross section through a centriole in the cortex of a protozoan. Each centriole is composed of nine sets of triplet microtubules arranged to form a cylinder. (C) Each triplet contains one complete microtubule (the A microtubule) fused to two incomplete microtubules (the B and C microtubules).


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Cell_Biology_Alberts. (D) The centriolar protein SAS-6 forms a coiled-coil dimer. Nine SAS-6 dimers can self-associate to form a ring. Located at the hub of the centriole cartwheel structure, the SAS-6 ring is thought to generate the ninefold symmetry of the centriole. (A, from from M. Paintrand, et al. J. Struct. Biol. 108:107, 1992. With permission from Elsevier; B, courtesy of Richard Linck; D, courtesy of Michel Steinmetz.) apical plasma membrane. From this asymmetrical location, a microtubule array extends along the long axis of the cell, with plus ends directed toward the basal surface (see Figure 16–4).


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Cell_Biology_Alberts. Microtubule polymerization dynamics are very different in cells than in solutions of pure tubulin. Microtubules in cells exhibit a much higher polymerization rate (typically 10–15 μm/min, relative to about 1.5 μm/min with purified tubulin at similar concentrations), a greater catastrophe frequency, and extended pauses in microtubule growth, a dynamic behavior rarely observed in pure tubulin solutions. These and other differences arise because microtubule dynamics inside the cell are governed by a variety of proteins that bind tubulin dimers or microtubules, as summarized in Panel 16–4.


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Cell_Biology_Alberts. Proteins that bind to microtubules are collectively called microtubule-associated proteins, or MAPs. Some MAPs can stabilize microtubules against disassembly. A subset of MAPs can also mediate the interaction of microtubules with other cell components. This subset is prominent in neurons, where stabilized micro-tubule bundles form the core of the axons and dendrites that extend from the cell body (Figure 16–50). These MAPs have at least one domain that binds to the microtubule surface and another that projects outward. The length of the projecting domain can determine how closely MAP-coated microtubules pack together, as demonstrated in cells engineered to overproduce different MAPs. Cells over-expressing MAP2, which has a long projecting domain, form bundles of stable microtubules that are kept widely spaced, while cells overexpressing tau, a MAP with a much shorter projecting domain, form bundles of more closely packed microtubules (Figure 16–51). MAPs are the targets of several


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Cell_Biology_Alberts. are kept widely spaced, while cells overexpressing tau, a MAP with a much shorter projecting domain, form bundles of more closely packed microtubules (Figure 16–51). MAPs are the targets of several protein kinases, and phosphorylation of a MAP can control both its activity and localization inside cells.


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Cell_Biology_Alberts. Cells contain numerous proteins that bind the ends of microtubules and thereby influence microtubule stability and dynamics. These proteins can influence the Figure 16–50 Localization of MAPs in the axon and dendrites of a neuron. This immunofluorescence micrograph shows the distribution of the proteins tau (green) and MAP2 (orange) in a hippocampal neuron in culture. Whereas tau staining is confined to the axon (long and branched in this neuron), MAP2 staining is confined to the cell body and its dendrites. The antibody used here to detect tau binds only to unphosphorylated tau; phosphorylated tau is also present in dendrites. (Courtesy of James W. Mandell and Gary A. Banker.) Figure 16–49 A microtubule array can find the center of a cell. After the arm of a fish pigment cell is cut off with a needle, the microtubules in the detached cell fragment reorganize so that their minus ends end up near the center of the fragment, buried in a new microtubule-organizing center.


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Cell_Biology_Alberts. nucleates assembly and remains associated with centrosome the minus end remain associated with growing plus ends prevents assembly and can link them to other structures, such as membranes binds subunits, links to intermediate flaments Some of the major accessory proteins of the microtubule cytoskeleton. Except for two classes of motor proteins, an example of each major type is shown. Each of these is discussed in the text. However, most cells contain more than a hundred different microtubule-binding proteins, and — as for the actin-associated proteins — it is likely that there are important types of microtubule-associated proteins that are not yet recognized.


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Cell_Biology_Alberts. rate at which a microtubule switches from a growing to a shrinking state (the frequency of catastrophes) or from a shrinking to a growing state (the frequency of rescues). For example, members of a family of kinesin-related proteins known as catastrophe factors (or kinesin-13) bind to microtubule ends and appear to pry protofilaments apart, lowering the normal activation-energy barrier that prevents a microtubule from springing apart into the curved protofilaments that are characteristic of the shrinking state (Figure 16–52). Another protein, called Nezha or Patronin, protects microtubule minus ends from the effects of catastrophe factors.


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Cell_Biology_Alberts. While very few microtubule minus-end-binding proteins have been characterized, a large subset of MAPs has been identified that are enriched at microtubule plus ends. A particularly ubiquitous example is XMAP215, which has close homologs in organisms that range from yeast to humans. XMAP215 binds free tubulin subunits and delivers them to the plus end, thereby promoting microtubule polymerization and simultaneously counteracting catastrophe factor activity (see Figure 16–52). The phosphorylation of XMAP215 during mitosis inhibits GTP cap on plus end of microtubule Figure 16–51 Organization of microtubule bundles by MAPs. (A) MAP2 binds along the microtubule lattice at one of its ends and extends a long projecting arm with a second microtubule-binding domain at the other end. (B) Tau possesses a shorter microtubule cross-linking domain.


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Cell_Biology_Alberts. Electron micrograph showing a cross section through a microtubule bundle in a cell overexpressing MAP2. The regular spacing of the microtubules (MTs) in this bundle results from the constant length of the projecting arms of the MAP2. Similar cross section through a microtubule bundle in a cell overexpressing tau. Here the microtubules are spaced more closely together than they are in (C) because of tau’s relatively short projecting arm. (C and D, courtesy of J. Chen et al., Nature 360:674–677, 1992. With permission from Macmillan Publishers Ltd.)


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Cell_Biology_Alberts. Figure 16–52 The effects of proteins that bind to microtubule ends. The transition between microtubule growth and shrinkage is controlled in cells by a variety of proteins. Catastrophe factors such as kinesin-13, a member of the kinesin motor protein superfamily, bind to microtubule ends and pry them apart, thereby promoting depolymerization. On the other hand, a MAP such as XMAP215 stabilizes the end of a growing microtubule (XMAP stands for Xenopus microtubuleassociated protein, and the number refers to its molecular mass in kilodaltons). XMAP215 binds tubulin dimers and delivers them to the microtubule plus end, thereby increasing the microtubule growth rate and suppressing catastrophes. its activity and shifts the balance of its competition with catastrophe factors. This shift results in a tenfold increase in the dynamic instability of microtubules during mitosis, a transition that is critical for the efficient construction of the mitotic spindle (discussed in Chapter 17).


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Cell_Biology_Alberts. In many cells, the minus ends of microtubules are stabilized by association with a capping protein or the centrosome, or else they serve as microtubule depolymerization sites. The plus ends, in contrast, efficiently explore and probe the entire cell space. Microtubule-associated proteins called plus-end tracking proteins (+TIPs) accumulate at these active ends and appear to rocket around the cell as passengers at the ends of rapidly growing microtubules, dissociating from the ends when the microtubules begin to shrink (Figure 16–53).


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Cell_Biology_Alberts. The kinesin-related catastrophe factors and XMAP215 mentioned above behave as +TIPs and act to modulate the growth and shrinkage of the microtubule end to which they are attached. Other +TIPs control microtubule positioning by helping to capture and stabilize the growing microtubule end at specific cellular targets, such as the cell cortex or the kinetochore of a mitotic chromosome. EB1 and its relatives, small dimeric proteins that are highly conserved in animals, plants, and fungi, are key players in this process. EB1 proteins do not actively move toward plus ends, but rather recognize a structural feature of the growing plus end (see Figure 16–53). Several of the +TIPs depend on EB1 proteins for their plus-end accumulation and also interact with each other and with the microtubule lattice. By attaching to the plus end, these factors allow the cell to harness the energy of microtubule polymerization to generate pushing forces that can be used for positioning the spindle,


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Cell_Biology_Alberts. lattice. By attaching to the plus end, these factors allow the cell to harness the energy of microtubule polymerization to generate pushing forces that can be used for positioning the spindle, chromosomes, or organelles.


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Cell_Biology_Alberts. As it does with actin monomers, the cell sequesters unpolymerized tubulin subunits to maintain a pool of active subunits at a level near the critical concentration. One molecule of the small protein stathmin (also called Op18) binds to two tubulin heterodimers and prevents their addition to the ends of microtubules (Figure 16–54). Stathmin thus decreases the effective concentration of tubulin subunits that are available for polymerization (an action analogous to that of the drug colchicine), and enhances the likelihood that a growing microtubule will switch to the shrinking state. Phosphorylation of stathmin inhibits its binding to tubulin, and signals that cause stathmin phosphorylation can increase the rate of microtubule elongation and suppress dynamic instability. Stathmin has been implicated in the regulation of both cell proliferation and cell death. Interestingly, mice lacking stathmin develop normally but are less fearful than wild-type mice, reflecting a role for stathmin in


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Cell_Biology_Alberts. in the regulation of both cell proliferation and cell death. Interestingly, mice lacking stathmin develop normally but are less fearful than wild-type mice, reflecting a role for stathmin in neurons of the amygdala, where it is normally expressed at high levels.


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Cell_Biology_Alberts. Severing is another mechanism employed by the cell to destabilize microtubules. To sever a microtubule, thirteen longitudinal bonds must be broken, one for each protofilament. The protein katanin, named after the Japanese word for Figure 16–53 +TIP proteins found at the growing plus ends of microtubules.


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Cell_Biology_Alberts. Figure 16–53 +TIP proteins found at the growing plus ends of microtubules. (A) Frames from a fluorescence time-lapse movie of the edge of a cell expressing fluorescently labeled tubulin that incorporates into microtubules (red) as well as the +TIP protein EB1 tagged with a different color (green). The same microtubule is marked (asterisk) in successive movie frames. When the microtubule is growing (frames 1, 2), EB1 is associated with the tip. When the microtubule undergoes a catastrophe and begins shrinking, EB1 is lost (frames 3, 4). The labeled EB1 is regained when growth of the microtubule is rescued (frame 5). See Movie 16.8. (B) In the fission yeast Schizosaccharomyces pombe, the plus ends of the microtubules (green) are associated with the homolog of EB1 (red) at the two poles of the rod-shaped cells. (A, courtesy of Anna Akhmanova and Ilya Grigoriev; B, courtesy of Takeshi Toda.) 2.5 nm


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Cell_Biology_Alberts. Figure 16–54 Sequestration of tubulin by stathmin. Structural studies with electron microscopy and crystallography suggest that the elongated stathmin protein binds along the side of two tubulin heterodimers. (Adapted from M.O. Steinmetz et al., EMBO J. 19:572–580, 2000. With permission from John Wiley and Sons.) “sword,” accomplishes this demanding task (Figure 16–55). Katanin is made up of two subunits: a smaller subunit that hydrolyzes ATP and performs the actual severing, and a larger one that directs katanin to the centrosome. Katanin releases microtubules from their attachment to a microtubule-organizing center and is thought to contribute to the rapid microtubule depolymerization observed at the poles of spindles during mitosis. It may also be involved in microtubule release and depolymerization in proliferating cells in interphase and in postmitotic cells such as neurons. Two Types of Motor Proteins Move Along Microtubules


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Cell_Biology_Alberts. Like actin filaments, microtubules also use motor proteins to transport cargo and perform a variety of other functions within the cell. There are two major classes of microtubule-based motors, kinesins and dyneins. Kinesin-1, also called “conventional kinesin,” was first purified from squid neurons, where it carries membrane-enclosed organelles away from the cell body toward the axon terminal by walking toward the plus end of microtubules. Kinesin-1 is similar to myosin II in having two heavy chains per active motor; these form two globular head motor domains that are held together by an elongated coiled-coil tail that is responsible for heavy-chain dimerization. One kinesin-1 light chain associates with each heavy chain through its tail domain and mediates cargo binding. Like myosin, kinesin is a member of a large protein superfamily, for which the motor domain is the common element (Figure 16–56). The yeast Saccharomyces cerevisiae has six distinct kinesins. The nematode C. elegans


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Cell_Biology_Alberts. is a member of a large protein superfamily, for which the motor domain is the common element (Figure 16–56). The yeast Saccharomyces cerevisiae has six distinct kinesins. The nematode C. elegans has 20 kinesins, and humans have 45.


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Cell_Biology_Alberts. There are at least fourteen distinct families in the kinesin superfamily. Most of them have the motor domain at the N-terminus of the heavy chain and walk toward the plus end of the microtubule. One family has the motor domain at the C-terminus and walks in the opposite direction, toward the minus end of the microtubule, while kinesin-13 has a central motor domain and does not walk at all, but uses the energy of ATP hydrolysis to depolymerize microtubule ends, as described above (see Figure 16–52). Some kinesin heavy chains are homodimers, and others are heterodimers. Most kinesins have a binding site in the tail for another microtubule; alternatively, they may link the motor to a membrane-enclosed organelle via a light chain or an adaptor protein. Many of the kinesin superfamily members have specific roles in mitotic spindle formation and in chromosome segregation during cell division.


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Cell_Biology_Alberts. In kinesin-1, instead of the rocking of a lever arm, small movements at the nucleotide-binding site regulate the docking and undocking of the motor head domain to a long linker region. This acts to throw the second head forward along Figure 16–55 Microtubule severing by katanin. Taxol stabilized, rhodaminelabeled microtubules were adsorbed on the surface of a glass slide, and purified katanin was added along with ATP. (A) There are a few breaks in the microtubules 30 seconds after the addition of katanin. (B) The same field 3 minutes after the addition of katanin. The filaments have been severed in many places, leaving a series of small fragments at the previous locations of the long microtubules. (From J.J. Hartman et al., Cell 93:277–287, 1998. With permission from Elsevier.)


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Cell_Biology_Alberts. Figure 16–56 Kinesin and kinesinrelated proteins. Structures of four kinesin superfamily members. As in the myosin superfamily, only the motor domains are conserved. Kinesin-1 has the motor domain at the N-terminus of the heavy chain. The middle domain forms a long coiled-coil, mediating dimerization. The C-terminal domain forms a tail that attaches to cargo, such as a membrane-enclosed organelle. Kinesin-5 forms tetramers where two dimers associate by their tails. The bipolar kinesin-5 tetramer is able to slide two microtubules past each other, analogous to the activity of the bipolar thick filaments formed by myosin II. Kinesin-13 has its motor domain located in the middle of the heavy chain. It is a member of a family of kinesins that have lost typical motor activity and instead bind to microtubule ends to promote depolymerization (see Figure 16–52). Kinesin-14 is a C-terminal kinesin that includes the Drosophila protein Ncd and the yeast protein Kar3. These kinesins generally


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Cell_Biology_Alberts. to microtubule ends to promote depolymerization (see Figure 16–52). Kinesin-14 is a C-terminal kinesin that includes the Drosophila protein Ncd and the yeast protein Kar3. These kinesins generally travel in the opposite direction from the majority of kinesins, toward the minus end instead of the plus end of a microtubule.


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Cell_Biology_Alberts. Figure 16–57 The mechanochemical cycle of kinesin. Kinesin-1 is a dimer of two nucleotide-binding motor domains (heads) that are connected through a long coiled-coil tail (see Figure 16–56). The two kinesin motor domains work in a coordinated manner; during a kinesin “step,” the rear head detaches from its tubulin binding site, passes the partner motor domain, and then rebinds to the next available tubulin binding site. Using this “handover-hand” motion, the kinesin dimer can move for long distances on the microtubule without completely letting go of its track.


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Cell_Biology_Alberts. At the start of each step, one of the two kinesin motor domain heads, the rear or lagging head (dark green), is tightly bound to the microtubule and to ATP, while the front or leading head is loosely bound to the microtubule with ADP in its binding site. The forward displacement of the rear motor domain is driven by the dissociation of ADP and binding of ATP in the leading head (between panels 2 and 3 in this drawing). The binding of ATP to this motor domain causes a small peptide called the “neck linker” to shift from a rearward-pointing to a forward-pointing conformation (the neck linker is drawn here as a purple connecting line between the leading motor domain and the intertwined coiled-coil). This shift pulls the rear head forward, once it has detached from the microtubule with ADP bound [detachment requires ATP hydrolysis and phosphate (Pi) release]. The kinesin molecule is now poised for the next step, which proceeds by an exact repeat of the process shown (Movie 16.9).


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Cell_Biology_Alberts. the protofilament to a binding site 8 nm closer to the microtubule plus end, which is the distance between tubulin dimers of a protofilament. The nucleotide-hydrolysis cycles in the two heads are closely coordinated, so that this cycle of linker docking and undocking allows the two-headed motor to move in a hand-overhand (or head-over-head) stepwise manner (Figure 16–57).


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Cell_Biology_Alberts. The dyneins are a family of minus-end directed microtubule motors unrelated to the kinesins. They are composed of one, two, or three heavy chains (that include the motor domain) and a large and variable number of associated intermediate, light-intermediate, and light chains. The dynein family has two major branches (Figure 16–58). The first branch contains the cytoplasmic dyneins, which are homodimers of two heavy chains. Cytoplasmic dynein 1 is encoded by a single gene in almost all eukaryotic cells, but is missing from flowering plants and some algae. It is used for organelle and mRNA trafficking, for positioning the centrosome and nucleus during cell migration, and for construction of the microtubule spindle in mitosis and meiosis. Cytoplasmic dynein 2 is found only in eukaryotic organisms that have cilia and is used to transport material from the tip to the base of the cilia, a process called intraflagellar transport. Axonemal dyneins (also called ciliary dyneins) comprise the


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Cell_Biology_Alberts. that have cilia and is used to transport material from the tip to the base of the cilia, a process called intraflagellar transport. Axonemal dyneins (also called ciliary dyneins) comprise the second branch and include monomers, heterodimers, and heterotrimers, with one, two, or three motor-containing heavy chains, respectively. They are highly specialized for the rapid and efficient sliding movements of microtubules that drive the beating of cilia and flagella (discussed later).


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Cell_Biology_Alberts. Figure 16–58 Dyneins. (A) Freeze-etch electron micrographs of a molecule of cytoplasmic dynein and a molecule of ciliary (axonemal) dynein. Like myosin II and kinesin-1, cytoplasmic dynein is a two-headed molecule. The ciliary dynein shown is from a protozoan and has three heads; ciliary dynein from animals has two heads. Note that the dynein head is very large compared with the head of either myosin or kinesin. (B) Schematic depiction of cytoplasmic dynein showing the two heavy chains (blue and gray) that contain domains for microtubule (MT) binding and ATP hydrolysis, connected by a long stalk. Bound to the heavy chain are multiple intermediate chains (dark green) and light chains (light green) that help to mediate many of dynein’s functions. (A, courtesy of John Heuser; B, adapted from R. Vale, Cell 112:467–480, 2003. With permission from Cell Press.) (B) power stroke, 8 nm


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Cell_Biology_Alberts. Dyneins are the largest of the known molecular motors, and they are also among the fastest: axonemal dyneins attached to a glass slide can move micro-tubules at the rate of 14 μm/sec. The dynein motor is structurally unrelated to myosins and kinesins, but still follows the general rule of coupling nucleotide hydrolysis to microtubule binding and unbinding as well as to a force-generating conformational change (Figure 16–59).


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Cell_Biology_Alberts. A major function of cytoskeletal motors in interphase cells is the transport and positioning of membrane-enclosed organelles (Movie 16.10). Kinesin was originally identified as the protein responsible for fast anterograde axonal transport, the rapid movement of mitochondria, secretory vesicle precursors, and various synapse components down the microtubule highways of the axon to the distant nerve terminals. Cytoplasmic dynein was identified as the motor responsible for transport in the opposite direction, retrograde axonal transport. Although organelles in most cells need not cover such long distances, their polarized transport is equally necessary. A typical microtubule array in an interphase cell is oriented with the minus ends near the center of the cell at the centrosome and the plus ends extending to the cell periphery. Thus, centripetal movements of organelles or vesicles toward the cell center require the action of minus-end directed cytoplasmic dynein motors, whereas


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Cell_Biology_Alberts. the plus ends extending to the cell periphery. Thus, centripetal movements of organelles or vesicles toward the cell center require the action of minus-end directed cytoplasmic dynein motors, whereas centrifugal movements toward the periphery require plus-end directed kinesin motors. Interestingly, in animal cells, nearly all minus-end directed transport is driven by the single cytoplasmic dynein 1 motor, whereas 15 different kinesins are used for plus-end directed transport.


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Cell_Biology_Alberts. Figure 16–59 The power stroke of dynein. (A) The organization of the domains in each dynein heavy chain. This is a huge polypeptide, containing nearly 4000 amino acids. The number of heavy chains in a dynein is equal to its number of motor heads. (B) Illustration of dynein c, a monomeric axonemal dynein found in the unicellular green alga Chlamydomonas reinhardtii. The large dynein motor head is a planar ring containing a C-terminal domain (gray) and six AAA domains, four of which retain ATP-binding sequences, but only one of which (dark red) has the major ATPase activity. Extending from the head are a long, coiled-coil stalk with the microtubule-binding site at the tip, and a tail that attaches to an adjacent microtubule in the axoneme. In the ATP-bound state, the stalk is detached from the microtubule, but ATP hydrolysis causes stalk–microtubule attachment (left). Subsequent release of ADP and phosphate (Pi) then leads to a large conformational “power stroke” involving rotation of


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Cell_Biology_Alberts. the microtubule, but ATP hydrolysis causes stalk–microtubule attachment (left). Subsequent release of ADP and phosphate (Pi) then leads to a large conformational “power stroke” involving rotation of the head and stalk relative to the tail (right). Each cycle generates a step of about 8 nm, thereby contributing to flagellar beating (see Figure 16–65). In the case of cytoplasmic dynein, the tail is attached to a cargo such as a vesicle, and a single power stroke transports the cargo about 8-nm along the microtubule toward its minus end (see Figure 16–60). (C) Electron micrographs of purified monomeric dyneins in two different conformations representing different steps in the mechanochemical cycle. (C, from


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Cell_Biology_Alberts. S.A. Burgess et al., Nature 421:715–718, 2003. With permission from Macmillan Publishers Ltd.)


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Cell_Biology_Alberts. A clear example of the effect of microtubules and microtubule motors on the behavior of intracellular membranes is their role in organizing the endoplasmic reticulum (ER) and the Golgi apparatus. The network of ER membrane tubules aligns with microtubules and extends almost to the edge of the cell (Movie 16.11), whereas the Golgi apparatus is located near the centrosome. When cells are treated with a drug that depolymerizes microtubules, such as colchicine or nocodazole, the ER collapses to the center of the cell, while the Golgi apparatus fragments and disperses throughout the cytoplasm. In vitro, kinesins can tether ER-derived membranes to preformed microtubule tracks and walk toward the microtubule plus ends, dragging the ER membranes out into tubular protrusions and forming a membranous web that looks very much like the ER in cells. Conversely, dyneins are required for positioning the Golgi apparatus near the cell center of animal cells; they do this by moving Golgi vesicles along


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Cell_Biology_Alberts. web that looks very much like the ER in cells. Conversely, dyneins are required for positioning the Golgi apparatus near the cell center of animal cells; they do this by moving Golgi vesicles along microtubule tracks toward the microtubules’ minus ends at the centrosome.


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Cell_Biology_Alberts. The different tails and their associated light chains on specific motor proteins allow the motors to attach to their appropriate organelle cargo. Membrane-associated motor receptors that are sorted to specific membrane-enclosed compartments interact directly or indirectly with the tails of the appropriate kinesin family members. Many viruses take advantage of microtubule motor-based transport during infection and use kinesin to move from their site of replication and assembly to the plasma membrane, from which they are poised to infect neighboring cells. An outer-membrane protein of Vaccinia virus, for example, contains an amino acid motif that mediates binding to kinesin-1 light chain and transport along microtubules to the plasma membrane. Interestingly, this motif is present in over 450 human proteins, one-third of which are associated with human diseases. Thus, kinesin transports a diverse set of cargoes involved in a wide range of important cellular functions.


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Cell_Biology_Alberts. For dynein, a large macromolecular assembly often mediates attachment to membranes. Cytoplasmic dynein, itself a huge protein complex, requires association with a second large protein complex called dynactin to translocate organelles effectively. The dynactin complex includes a short, actin-like filament that forms from the actin-related protein Arp1 (distinct from Arp2 and Arp3, the components of the Arp 2/3 complex involved in the nucleation of conventional actin filaments) (Figure 16–60). A number of other proteins also contribute to dynein cargo binding and motor regulation, and their function is especially important in neurons, where defects in microtubule-based transport have been linked to neurological diseases. A striking example is smooth brain, or lissencephaly, a human disorder in which cells fail to migrate to the cerebral cortex of the developing brain. One type of lissencephaly is caused by defects in Lis1, a dynein-binding protein required for nuclear migration in


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Cell_Biology_Alberts. in which cells fail to migrate to the cerebral cortex of the developing brain. One type of lissencephaly is caused by defects in Lis1, a dynein-binding protein required for nuclear migration in several species. In the normal brain, migration of the nucleus directs the developing neural cell body toward its correct position in the cortex. In the absence of Lis1, however, the nuclei of migrating neurons fail to attach to dynein, resulting in nuclear-migration defects. Dynein is required continuously for neuronal function, as mutations in a dynactin subunit or in the tail region of cytoplasmic dynein lead to neuronal degeneration in humans and mice. These effects are associated with decreased retrograde axonal transport and provide strong evidence for the importance of robust axonal transport in neuronal viability.


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Cell_Biology_Alberts. The cell can regulate the activity of motor proteins and thereby cause either a change in the positioning of its membrane-enclosed organelles or whole-cell movements. Fish melanocytes provide one of the most dramatic examples. These giant cells, which are responsible for rapid changes in skin coloration in several species of fish, contain large pigment granules that can alter their location in response to neuronal or hormonal stimulation (Figure 16–61). The pigment granules aggregate or disperse by moving along an extensive network of microtubules that are anchored at the centrosome by their minus ends. The tracking of individual pigment granules reveals that the inward movement is rapid and smooth, while the outward movement is jerky, with frequent backward steps. Both dynein and kinesin microtubule motors are associated with the pigment granules. The


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Cell_Biology_Alberts. Figure 16–60 Dynactin mediates the attachment of dynein to a membrane-enclosed organelle. Dynein requires the presence of a large number of accessory proteins to associate with membrane-enclosed organelles. Dynactin is a large complex that includes components that bind weakly to microtubules, components that bind to dynein itself, and components that form a small, actin-like filament made of the actin-related protein Arp1. jerky outward movements may result from a tug-of-war between the two opposing microtubule motor proteins, with the stronger kinesin winning out overall. When intracellular cyclic AMP levels decrease, kinesin is inactivated, leaving dynein free to drag the pigment granules rapidly toward the cell center, changing the fish’s color. In a similar way, the movement of other membrane organelles coated with particular motor proteins is controlled by a complex balance of competing signals that regulate both motor protein attachment and activity.


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Cell_Biology_Alberts. Construction of Complex Microtubule Assemblies Requires Microtubule Dynamics and Motor Proteins The construction of the mitotic spindle and the neuronal cytoskeleton are important and fascinating examples of the power of organization by teams of motor proteins interacting with dynamic cytoskeletal filaments. As described in Chapter 17, mitotic spindle assembly depends on reorganization of the interphase array of microtubules to form a bipolar array of microtubules, with their minus ends focused at the poles and their plus ends overlapping in the center or connecting to chromosomes. Spindle assembly depends on the coordinated actions of several motor proteins and other factors that modulate polymerization dynamics (see Figures 17–23 and 17–25).


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Cell_Biology_Alberts. Neurons also contain complex cytoskeletal structures. As they differentiate, neurons send out specialized processes that will either receive electrical signals (dendrites) or transmit electrical signals (axons) (see Figure 16–50). The beautiful and elaborate branching morphology of axons and dendrites enables neurons to form tremendously complex signaling networks, interacting with many other cells simultaneously and making possible the complicated behavior of the higher animals. Both axons and dendrites (collectively called neurites) are filled with bundles of microtubules that are critical to both their structure and their function. In axons, all the microtubules are oriented in the same direction, with their minus end pointing back toward the cell body and their plus end pointing toward the axon terminals (Figure 16–62). The microtubules do not reach from the cell Figure 16–61 Regulated melanosome movements in fish pigment cells.


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Cell_Biology_Alberts. Figure 16–61 Regulated melanosome movements in fish pigment cells. These giant cells, which are responsible for changes in skin coloration in several species of fish, contain large pigment granules, or melanosomes (brown). The melanosomes can change their location in the cell in response to a hormonal or neuronal stimulus. (A) Schematic view of a pigment cell, showing the dispersal and aggregation of melanosomes in response to an increase or decrease in intracellular cyclic AMP (cAMP), respectively. Both redistributions of melanosomes occur along microtubules. (B) Bright-field images of a single cell in a scale of an African cichlid fish, showing its melanosomes either dispersed throughout the cytoplasm (left) or aggregated in the center of the cell (right). (B, courtesy of Leah Haimo.)


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Cell_Biology_Alberts. Figure 16–62 Microtubule organization in a neuron. In a neuron, microtubule organization is complex. In the axon, all microtubules share the same polarity, with the plus ends pointing outward toward the axon terminus. No single microtubule stretches the entire length of the axon; instead, short overlapping segments of parallel microtubules make the tracks for fast axonal transport. In dendrites, the microtubules are of mixed polarity, with some plus ends pointing outward and some pointing inward. Vesicles can associate with both kinesin and dynein and move in either direction along the microtubules in axons and dendrites, depending on which motor is active.


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Cell_Biology_Alberts. body all the way to the axon terminals; each is typically only a few micrometers in length, but large numbers are staggered in an overlapping array. These aligned microtubule tracks act as a highway to transport specific proteins, protein-containing vesicles, and mRNAs to the axon terminals, where synapses are constructed and maintained. The longest axon in the human body reaches from the base of the spinal cord to the foot and is up to a meter in length. By comparison, dendrites are generally much shorter than axons. The microtubules in dendrites lie parallel to one another but their polarities are mixed, with some pointing their plus ends toward the dendrite tip, while others point back toward the cell body, reminiscent of the antiparallel microtubule array of the mitotic spindle.


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Cell_Biology_Alberts. Just as myofibrils are highly specialized and efficient motility machines built from actin and myosin filaments, cilia and flagella are highly specialized and efficient motility structures built from microtubules and dynein. Both cilia and flagella are hairlike cell appendages that have a bundle of microtubules at their core. Flagella are found on sperm and many protozoa. By their undulating motion, they enable the cells to which they are attached to swim through liquid media. Cilia are organized in a similar fashion, but they beat with a whiplike motion that resembles the breaststroke in swimming. Ciliary beating can either propel single cells through a fluid (as in the swimming of the protozoan Paramecium) or can move fluid over the surface of a group of cells in a tissue. In the human body, huge numbers of cilia (109/cm2 or more) line our respiratory tract, sweeping layers of mucus, trapped particles of dust, and bacteria up to the mouth where they are swallowed and ultimately


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Cell_Biology_Alberts. body, huge numbers of cilia (109/cm2 or more) line our respiratory tract, sweeping layers of mucus, trapped particles of dust, and bacteria up to the mouth where they are swallowed and ultimately eliminated. Likewise, cilia along the oviduct help to sweep eggs toward the uterus.


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Cell_Biology_Alberts. The movement of a cilium or a flagellum is produced by the bending of its core, which is called the axoneme. The axoneme is composed of microtubules and their associated proteins, arranged in a distinctive and regular pattern. Nine special doublet microtubules (comprising one complete and one partial micro-tubule fused together so that they share a common tubule wall) are arranged in a ring around a pair of single microtubules (Figure 16–63). Almost all forms of motile eukaryotic flagella and cilia (from protozoans to humans) have this characteristic arrangement. The microtubules extend continuously for the length of the axoneme, which can be 10–200 μm. At regular positions along the length of the microtubules, accessory proteins cross-link the microtubules together.


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Cell_Biology_Alberts. Figure 16–63 The arrangement of microtubules in a flagellum or cilium. (A) Electron micrograph of the flagellum of a green-alga cell (Chlamydomonas) shown in cross section, illustrating the distinctive “9 + 2” arrangement of microtubules. (B) Diagram of the parts of a flagellum or cilium. The various projections from the microtubules link the microtubules together and occur at regular intervals along the length of the axoneme. (C) High-resolution electron tomography image of an outer doublet microtubule showing structural details and features inside the microtubules called microtubule inner proteins (MIPs). (A, courtesy of Lewis Tilney; C, courtesy of Daniela Nicastro.) 942 Chapter 16: The Cytoskeleton


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Cell_Biology_Alberts. Molecules of axonemal dynein form bridges between the neighboring doublet microtubules around the circumference of the axoneme (Figure 16–64). When the motor domain of this dynein is activated, the dynein molecules attached to one microtubule doublet (see Figure 16–59) attempt to walk along the adjacent microtubule doublet, tending to force the adjacent doublets to slide relative to one another, much as actin thin filaments slide during muscle contraction. However, the presence of other links between the microtubule doublets prevents this sliding, and the dynein force is instead converted into a bending motion (Figure 16–65).


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Cell_Biology_Alberts. In humans, hereditary defects in axonemal dynein cause a condition called primary ciliary dyskinesia or Kartagener’s syndrome. This syndrome is characterized by inversion of the normal asymmetry of internal organs (sinus inversus) due to disruption of fluid flow in the developing embryo, male sterility due to immotile sperm, and a high susceptibility to lung infections due to paralyzed cilia being unable to clear the respiratory tract of debris and bacteria. Bacteria also swim using cell-surface structures called flagella, but these do not contain microtubules or dynein and do not wave or beat. Instead, bacterial flagella are long, rigid helical filaments, made up of repeating subunits of the protein flagellin. The flagella rotate like propellers, driven by a special rotary motor embedded in the bacterial cell wall. The use of the same name to denote these two very different types of swimming apparatus is an unfortunate historical accident.


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Cell_Biology_Alberts. Many cells possess a shorter, nonmotile counterpart of cilia and flagella called the primary cilium. Primary cilia can be viewed as specialized cellular compartments or organelles that perform a wide range of cellular functions, but share


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Cell_Biology_Alberts. Figure 16–64 Ciliary dynein. Ciliary (axonemal) dynein is a large protein assembly (nearly 2 million daltons) composed of 9–12 polypeptide chains, the largest of which is the heavy chain of more than 500,000 daltons. (A) The heavy chains form the major portion of the globular head and stem domains, and many of the smaller chains are clustered around the base of the stem. There are two heads in the outer dynein in metazoans (shown here), but three heads in protozoa, each formed from their own heavy chain. The tail of the molecule binds tightly to an A microtubule, while the large globular heads have an ATP-dependent binding site for a B microtubule (see Figure 16–63). When the heads hydrolyze their bound ATP, they move toward the minus end of the B microtubule, thereby producing a sliding force between the adjacent microtubule doublets in a cilium or flagellum (see Figure 16–59). (B) Freeze-etch electron micrograph of a cilium showing the dynein arms projecting at regular intervals


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Cell_Biology_Alberts. force between the adjacent microtubule doublets in a cilium or flagellum (see Figure 16–59). (B) Freeze-etch electron micrograph of a cilium showing the dynein arms projecting at regular intervals from the doublet microtubules. (B, courtesy of John Heuser.)


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Cell_Biology_Alberts. Figure 16–65 The bending of an axoneme. (A) When axonemes are exposed to the proteolytic enzyme trypsin, the linkages holding neighboring doublet microtubules together are broken. In this case, the addition of ATP allows the motor action of the dynein heads to slide one pair of doublet microtubules against the other pair. (B) In an intact axoneme (such as in a sperm), flexible protein links prevent the sliding of the doublet. The motor action therefore causes a bending motion, creating waves or beating motions.


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Cell_Biology_Alberts. many structural features with motile cilia. Both motile and nonmotile cilia are generated during interphase at plasma-membrane-associated structures called basal bodies that firmly root them at the cell surface. At the core of each basal body is a centriole, the same structure found embedded at the center of animal centrosomes, with nine groups of fused triplet microtubules arranged in a cartwheel (see Figure 16–48). Centrioles are multifunctional, contributing to assembly of the mitotic spindle in dividing cells but migrating to the plasma membrane of inter-phase cells to template the nucleation of the axoneme (Figure 16–66). Because no protein translation occurs in cilia, construction of the axoneme requires intraflagellar transport (IFT), a transport system discovered in the green algae Chlamydomonas. Analogous to the axon, motors move cargoes in both anterograde and retrograde directions, in this case driven by kinesin-2 and cytoplasmic dynein 2, respectively.


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Cell_Biology_Alberts. Primary cilia are found on the surface of almost all cell types, where they sense and respond to the exterior environment, functions best understood in the context of smell and sight. In the nasal epithelium, cilia protruding from dendrites of olfactory neurons are the site of both odorant reception and signal amplification. Similarly, the rod and cone cells of the vertebrate retina possess a primary cilium equipped with an expanded tip called the outer segment, which is specialized for converting light into a neural signal (see Figure 15–38). Maintenance of the outer segment requires continuous IFT-mediated transport of large quantities of lipids and proteins into the cilium, at rates of up to 2000 molecules per minute. The links between cilia function and the senses of sight and smell are underscored by Bardet-Biedl syndrome, a set of disorders associated with defects in IFT, the cilium, or the basal body. Patients with Bardet-Biedl syndrome cannot smell and suffer from retinal


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Cell_Biology_Alberts. are underscored by Bardet-Biedl syndrome, a set of disorders associated with defects in IFT, the cilium, or the basal body. Patients with Bardet-Biedl syndrome cannot smell and suffer from retinal degeneration. Other characteristics of this multifaceted disorder include hearing loss, polycystic kidney disease, diabetes, obesity, and polydactyly, suggesting that primary cilia have functions in many aspects of human physiology.


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Cell_Biology_Alberts. Microtubules are stiff polymers of tubulin molecules. They assemble by addition of GTP-containing tubulin subunits to the free end of a microtubule, with one end (the plus end) growing faster than the other. Hydrolysis of the bound GTP takes place Figure 16–66 Primary cilia. (A) Electron micrograph and diagram of the basal body of a mouse neuron primary cilium. The axoneme of the primary cilium (black arrow) is nucleated by the mother centriole at the basal body, which localizes at the plasma membrane near the cell surface.


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Cell_Biology_Alberts. (B) Centrioles function alternately as basal bodies and as the core of centrosomes. Before a cell enters the cell division cycle, the primary cilium is shed or resorbed. The centrioles recruit pericentriolar material and duplicate during S phase, generating two centrosomes, each of which contains a pair of centrioles. The centrosomes nucleate microtubules and localize to the poles of the mitotic spindle. Upon exit from mitosis, a primary cilium again grows from the mother centriole. (A, courtesy of Josef Spacek.) after assembly and weakens the bonds that hold the microtubule together. Microtubules are dynamically unstable and liable to catastrophic disassembly, but they can be stabilized in cells by association with other structures. Microtubule-organizing centers such as centrosomes protect the minus ends of microtubules and continually nucleate the formation of new microtubules. Microtubule-associated proteins (MAPs) stabilize microtubules, and those that localize to the plus end


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Cell_Biology_Alberts. the minus ends of microtubules and continually nucleate the formation of new microtubules. Microtubule-associated proteins (MAPs) stabilize microtubules, and those that localize to the plus end (+TIPs) can alter the dynamic properties of the microtubule or mediate their interaction with other structures. Counteracting the stabilizing activity of MAPs are catastrophe factors, such as kinesin-13 proteins, that act to peel apart microtubule ends. Other kinesin family members as well as dynein use the energy of ATP hydrolysis to move unidirectionally along a microtubule. The motor dynein moves toward the minus end of microtubules, and its sliding of axonemal microtubules underlies the beating of cilia and flagella. Primary cilia are nonmotile sensory organs found on many cell types.


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Cell_Biology_Alberts. All eukaryotic cells contain actin and tubulin. But the third major type of cytoskeletal protein, the intermediate filament, forms a cytoplasmic filament only in some metazoans—including vertebrates, nematodes, and mollusks. Intermediate filaments are particularly prominent in the cytoplasm of cells that are subject to mechanical stress and are generally not found in animals that have rigid exoskeletons, such as arthropods and echinoderms. It seems that intermediate filaments impart mechanical strength to tissues for the squishier animals.


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Cell_Biology_Alberts. Cytoplasmic intermediate filaments are closely related to their ancestors, the much more prevalent nuclear lamins, which are found in many eukaryotes but missing from unicellular organisms. The nuclear lamins form a meshwork lining the inner membrane of the nuclear envelope, where they provide anchorage sites for chromosomes and nuclear pores. Several times during metazoan evolution, lamin genes have apparently duplicated, and the duplicates have evolved to produce ropelike, cytoplasmic intermediate filaments. In contrast to the highly conserved actins and tubulin isoforms that are encoded by a handful of genes, different families of intermediate filaments are much more diverse and are encoded by 70 different human genes with distinct, cell type-specific functions (Table 16–2). (C) 0.1 µm NH2 COOH COOH NH2 staggered tetramer of two coiled-coil dimers lateral association of 8 tetramers addition of 8 tetramers to growing flament


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Cell_Biology_Alberts. (C) 0.1 µm NH2 COOH COOH NH2 staggered tetramer of two coiled-coil dimers lateral association of 8 tetramers addition of 8 tetramers to growing flament Figure 16–67 A model of intermediate filament construction. The monomer shown in (A) pairs with another monomer to form a dimer (B), in which the conserved central rod domains are aligned in parallel and wound together into a coiled-coil.


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Cell_Biology_Alberts. (C) Two dimers then line up side by side to form an antiparallel tetramer of four polypeptide chains. Dimers and tetramers are the soluble subunits of intermediate filaments. (D) Within each tetramer, the two dimers are offset with respect to one another, thereby allowing it to associate with another tetramer. (E) In the final 10-nm ropelike filament, tetramers are packed together in a helical array, which has 16 dimers (32 coiled-coils) in cross section. Half of these dimers are pointing in each direction. An electron micrograph of intermediate filaments is shown on the upper left (Movie 16.12). (Electron micrograph courtesy of Roy Quinlan.) Intermediate Filament Structure Depends on the Lateral Bundling and Twisting of Coiled-Coils


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Cell_Biology_Alberts. Although their aminoand carboxy-terminal domains differ, all intermediate filament family members are elongated proteins with a conserved central α-helical domain containing 40 or so heptad repeat motifs that form an extended coiled-coil structure with another monomer (see Figure 3–9). A pair of parallel dimers then associates in an antiparallel fashion to form a staggered tetramer (Figure 16–67). Unlike actin or tubulin subunits, intermediate filament subunits do not contain a binding site for a nucleotide. Furthermore, since the tetrameric subunit is made up of two dimers pointing in opposite directions, its two ends are the same. The assembled intermediate filament therefore lacks the overall structural polarity that is critical for actin filaments and microtubules. The tetramers pack together laterally to form the filament, which includes eight parallel protofilaments made up of tetramers. Each individual intermediate filament therefore has a cross section of 32 individual


Cell_Biology_Alberts

Cell_Biology_Alberts. pack together laterally to form the filament, which includes eight parallel protofilaments made up of tetramers. Each individual intermediate filament therefore has a cross section of 32 individual α-helical coils. This large number of polypeptides all lined up together, with the strong lateral hydrophobic interactions typical of coiled-coil proteins, gives intermediate filaments a ropelike character. They can be easily bent, with a persistence length of less than one micrometer (compared to several millimeters for microtubules and about ten micrometers for actin), but they are extremely difficult to break and can be stretched to over three times their length (see Figure 16–6).


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Cell_Biology_Alberts. Less is understood about the mechanism of assembly and disassembly of intermediate filaments than of actin filaments and microtubules. In pure protein solutions, intermediate filaments are extremely stable due to tight association of subunits, but some types of intermediate filaments, including vimentin, form highly dynamic structures in cells such as fibroblasts. Protein phosphorylation probably regulates their disassembly, in much the same way that phosphorylation regulates the disassembly of nuclear lamins in mitosis (see Figure 12–18). As evidence for rapid turnover, labeled subunits microinjected into tissue-culture cells incorporate into intermediate filaments within a few minutes. Remodeling of the intermediate filament network accompanies events requiring dynamic cellular reorganization, such as division, migration, and differentiation. Intermediate Filaments Impart Mechanical Stability to Animal Cells


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Cell_Biology_Alberts. Keratins are the most diverse intermediate filament family: there are about 20 found in different types of human epithelial cells and about 10 more that are specific to hair and nails; analysis of the human genome sequence has revealed that there are 54 distinct keratins. Every keratin filament is made up of an equal mixture of type I (acidic) and type II (neutral/basic) keratin proteins; these form a heterodimer filament subunit (see Figure 16–67). Cross-linked keratin networks held together by disulfide bonds can survive even the death of their cells, forming tough coverings for animals, as in the outer layer of skin and in hair, nails, claws, and scales. The diversity in keratins is clinically useful in the diagnosis of epithelial cancers (carcinomas), as the particular set of keratins expressed gives an indication of the epithelial tissue in which the cancer originated and thus can help to guide the choice of treatment.


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Cell_Biology_Alberts. A single epithelial cell may produce multiple types of keratins, and these copolymerize into a single network (Figure 16–68). Keratin filaments impart mechanical strength to epithelial tissues in part by anchoring the intermediate filaments at sites of cell–cell contact, called desmosomes, or cell–matrix contact, called hemidesmosomes (see Figure 16–4). We discuss these important adhesive structures in Chapter 19. Accessory proteins, such as filaggrin, bundle keratin filaments in differentiating cells of the epidermis to give the outermost layers of the


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Cell_Biology_Alberts. Figure 16–68 Keratin filaments in epithelial cells. Immunofluorescence micrograph of the network of keratin filaments (blue) in a sheet of epithelial cells in culture. The filaments in each cell are indirectly connected to those of its neighbors by desmosomes (discussed in Chapter 19). A second protein (red) has been stained to reveal the location of the cell boundaries. (Courtesy of Kathleen Green and Evangeline Amargo.) skin their special toughness. Individuals with mutations in the gene encoding filaggrin are strongly predisposed to dry skin diseases such as eczema.


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Cell_Biology_Alberts. Mutations in keratin genes cause several human genetic diseases. For example, when defective keratins are expressed in the basal cell layer of the epidermis, they produce a disorder called epidermolysis bullosa simplex, in which the skin blisters in response to even very slight mechanical stress, which ruptures the basal cells (Figure 16–69). Other types of blistering diseases, including disorders of the mouth, esophageal lining, and the cornea of the eye, are caused by mutations in the different keratins whose expression is specific to those tissues. All of these maladies are typified by cell rupture as a consequence of mechanical trauma and a disorganization or clumping of the keratin filament cytoskeleton. Many of the specific mutations that cause these diseases alter the ends of the central rod domain, demonstrating the importance of this particular part of the protein for correct filament assembly.


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Cell_Biology_Alberts. Members of another family of intermediate filaments, called neurofilaments, are found in high concentrations along the axons of vertebrate neurons (Figure 16–70). Three types of neurofilament proteins (NF-L, NF-M, and NF-H) coassemble in vivo, forming heteropolymers. The NF-H and NF-M proteins have lengthy C-terminal tail domains that bind to neighboring filaments, generating aligned arrays with a uniform interfilament spacing. During axonal growth, new neurofilament subunits are incorporated all along the axon in a dynamic process that involves the addition of subunits along the filament length as well as the ends. After an axon has grown and connected with its target cell, the diameter of the axon may increase as much as fivefold. The level of neurofilament gene expression seems to directly control axonal diameter, which in turn influences how fast electrical signals travel down the axon. In addition, neurofilaments provide strength and stability to the long cell processes of


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Cell_Biology_Alberts. directly control axonal diameter, which in turn influences how fast electrical signals travel down the axon. In addition, neurofilaments provide strength and stability to the long cell processes of neurons.


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Cell_Biology_Alberts. The neurodegenerative disease amyotrophic lateral sclerosis (ALS, or Lou Gehrig’s disease) is associated with an accumulation and abnormal assembly of neurofilaments in motor neuron cell bodies and in the axon, aberrations that may interfere with normal axonal transport. The degeneration of the axons leads to muscle weakness and atrophy, which is usually fatal. The overexpression of human NF-L or NF-H in mice results in mice that have an ALS-like disease. However, a causative link between neurofilament pathology and ALS has not been firmly established. basal cell of epidermis


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Cell_Biology_Alberts. basal cell of epidermis Figure 16–69 blistering of the skin caused by a mutant keratin gene. A mutant gene encoding a truncated keratin protein (lacking both the Nand C-terminal domains) was expressed in a transgenic mouse. The defective protein assembles with the normal keratins and thereby disrupts the keratin filament network in the basal cells of the skin. Light micrographs of cross sections of (A) normal and (B) mutant skin show that the blistering results from the rupturing of cells in the basal layer of the mutant epidermis (short red arrows). (C) A sketch of three cells in the basal layer of the mutant epidermis, as observed by electron microscopy. As indicated by the red arrow, the cells rupture between the nucleus and the hemidesmosomes (discussed in Chapter 19), which connect the keratin filaments to the underlying basal lamina. (From P.A. Coulombe et al., J. Cell Biol. 115:1661–1674, 1991. With permission from The Rockefeller University Press.)


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Cell_Biology_Alberts. The vimentin-like filaments are a third family of intermediate filaments. Desmin, a member of this family, is expressed in skeletal, cardiac, and smooth muscle, where it forms a scaffold around the Z disc of the sarcomere (see Figure 16–34). Mice lacking desmin show normal initial muscle development, but adults have various muscle-cell abnormalities, including misaligned muscle fibers. In humans, mutations in desmin are associated with various forms of muscular dystrophy and cardiac myopathy, illustrating the important role of desmin in stabilizing muscle fibers.


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Cell_Biology_Alberts. Besides their well-established role in maintaining the mechanical stability of the nucleus, it is becoming increasingly evident that one class of lamins, the A-type, together with many proteins of the nuclear envelope, are scaffolds for proteins that control myriad cellular processes including transcription, chromatin organization, and signal transduction. The majority of laminopathies are associated with mutant versions of lamin A and include tissue-specific diseases. Skeletal and cardiac abnormalities might be explained by a weakened nuclear envelope leading to cell damage and death, but laminopathies are also thought to arise from pathogenic and tissue-specific alterations in gene expression. Linker Proteins Connect Cytoskeletal Filaments and Bridge the Nuclear Envelope


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Cell_Biology_Alberts. Linker Proteins Connect Cytoskeletal Filaments and Bridge the Nuclear Envelope The intermediate filament network is linked to the rest of the cytoskeleton by members of a family of proteins called plakins. Plakins are large and modular, containing multiple domains that connect cytoskeletal filaments to each other and to junctional complexes. Plectin is a particularly interesting example. In addition to bundling intermediate filaments, it links the intermediate filaments to microtubules, actin filament bundles, and filaments of the motor protein myosin II; it also helps attach intermediate filament bundles to adhesive structures at the plasma membrane (Figure 16–71). Plectin and other plakins can interact with protein complexes that connect the cytoskeleton to the nuclear interior. These complexes consist of SUN proteins 0.5 µm Figure 16–70 Two types of intermediate filaments in cells of the nervous system.


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Cell_Biology_Alberts. Figure 16–70 Two types of intermediate filaments in cells of the nervous system. Freeze-etch electron microscopic image of neurofilaments in a nerve cell axon, showing the extensive cross-linking through protein cross-bridges—an arrangement believed to give this long cell process great tensile strength. The cross-bridges are formed by the long, nonhelical extensions at the C-terminus of the largest neurofilament protein (NF-H). (B) Freeze-etch image of glial filaments in glial cells, showing that these intermediate filaments are smooth and have few cross-bridges. Conventional transmission electron micrograph of a cross section of an axon showing the regular side-to-side spacing of the neurofilaments, which greatly outnumber the microtubules. (A and B, courtesy of Nobutaka Hirokawa; C, courtesy of John Hopkins.)


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Cell_Biology_Alberts. Figure 16–71 Plectin cross-linking of diverse cytoskeletal elements. Plectin (green) is seen here making cross-links from intermediate filaments (blue) to microtubules (red). In this electron micrograph, the dots (yellow) are gold particles linked to anti-plectin antibodies. The entire actin filament network was removed to reveal these proteins. (From


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Cell_Biology_Alberts. T.M. Svitkina et al., J. Cell Biol. 135:991– 1007, 1996. With permission from The Rockefeller University Press.) of the inner nuclear membrane and KASH proteins (also called nesprins) of the outer nuclear membrane (Figure 16–72). SUN and KASH proteins bind to each other within the lumen of the nuclear envelope, forming a bridge that connects the nuclear and cytoplasmic cytoskeletons. Inside the nucleus, the SUN proteins bind to the nuclear lamina or chromosomes, whereas in the cytoplasm, KASH proteins can bind directly to actin filaments and indirectly to microtubules and intermediate filaments through association with motor proteins and plakins, respectively. This linkage serves to mechanically couple the nucleus to the cytoskeleton and is involved in many cellular functions, including chromosome movements inside the nucleus during meiosis, nuclear and centrosome positioning, nuclear migration, and global cytoskeletal organization.


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Cell_Biology_Alberts. Mutations in the gene for plectin cause a devastating human disease that combines epidermolysis bullosa (caused by disruption of skin keratin filaments), muscular dystrophy (caused by disruption of desmin filaments), and neurodegeneration (caused by disruption of neurofilaments). Mice lacking a functional plectin gene die within a few days of birth, with blistered skin and abnormal skeletal and heart muscles. Thus, although plectin may not be necessary for the initial formation and assembly of intermediate filaments, its cross-linking action is required to provide cells with the strength they need to withstand the mechanical stresses inherent to vertebrate life.


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Cell_Biology_Alberts. GTP-binding proteins called septins serve as an additional filament system in all eukaryotes except terrestrial plants. Septins assemble into nonpolar filaments that form rings and cagelike structures, which act as scaffolds to compartmentalize membranes into distinct domains, or recruit and organize the actin and microtubule cytoskeletons. First identified in budding yeast, septin filaments localize to the neck between a dividing yeast mother cell and its growing bud (Figure 16–73A). At this location, septins block the movement of proteins from one side of the bud neck to the other, thereby concentrating cell growth preferentially within the bud. Septins also recruit the actin–myosin machinery that forms the contractile ring required for cytokinesis. In animal cells, septins function in cell division, migration, and vesicle trafficking. In primary cilia, for example, a ring of septin filaments assembles at the base of the cilium and serves as a diffusion barrier at the plasma


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Cell_Biology_Alberts. in cell division, migration, and vesicle trafficking. In primary cilia, for example, a ring of septin filaments assembles at the base of the cilium and serves as a diffusion barrier at the plasma membrane, restricting the movement of membrane proteins and establishing a specific composition in the ciliary membrane (Figure 16–73B and C). Reduction of septin levels impairs primary cilium formation and signaling.


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Cell_Biology_Alberts. There are 7 septin genes in yeast and 13 in human, and septin proteins fall into four groups on the basis of sequence relationships. In a test tube, purified septins assemble into symmetrical hetero-hexamers or hetero–octamers that Figure 16–72 SUN–KASH protein complexes connect the nucleus and cytoplasm through the nuclear envelope. The cytoplasmic cytoskeleton is linked across the nuclear envelope to the nuclear lamina or chromosomes through SUN and KASH proteins (orange and purple, respectively). The SUN and KASH domains of these proteins bind within the lumen of the nuclear envelope. From the inner nuclear envelope, SUN proteins connect to the nuclear lamina or chromosomes. KASH proteins in the outer nuclear envelope connect to the cytoplasmic cytoskeleton by binding microtubule motor proteins, actin filaments, or plectin.


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Cell_Biology_Alberts. mother cell 0.5 µm 10 µm2 µm form nonpolar paired filaments (Figure 16–74). GTP binding is required for the folding of septin polypeptides, but the role of GTP hydrolysis in septin function is not understood. Septin structures assemble and disassemble inside cells, but they are not as dynamic as actin filaments and microtubules. Whereas tubulin and actin have been highly conserved in evolution, intermediate filament proteins are very diverse. There are many tissue-specific forms of intermediate filaments in the cytoplasm of animal cells, including keratin filaments in epithelial cells, neurofilaments in nerve cells, and desmin filaments in muscle cells. The primary function of these filaments is to provide mechanical strength. Septins comprise an additional system of filaments that organize compartments inside cells. Figure 16–74 Septins polymerize to form paired filaments and sheets.


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Cell_Biology_Alberts. Figure 16–74 Septins polymerize to form paired filaments and sheets. (A) Electron micrograph of a septin rod assembled by combining two copies each of the four yeast septins illustrated at the right. The eight-subunit rod is nonpolar because the central pair of subunits (Cdc10) creates a symmetrical dimer. (B) Electron micrograph of paired septin filaments and sheets, assembled from purified septins in the presence of high salt concentrations. (C) Paired septin filaments may assemble by lateral association between filaments, mediated by coiled-coils formed between the paired C-terminal extensions of Cdc3 and Cdc12 that project from each filament. (Images and schematics adapted from A. Bertin et al., Proc. Natl Acad. Sci. USA 105:8274–8279, 2008. With permission from the National Academy of Sciences.)


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Cell_Biology_Alberts. Figure 16–73 Cell compartmentalization by septins. (A) Septins form filaments in the neck region between a mother yeast cell and bud. (B) In this photomicrograph of human cultured cells, the DNA is stained blue and septins are labeled in green. The microtubules of primary cilia are labeled with an antibody that recognizes a modified (acetylated) form of tubulin (red) that is enriched in the axoneme. (C) A magnified image reveals a collar of septin at the base of the cilium. (A, from B. Byers and L. Goetsch, J. Cell Biol. 69:717–721, 1976. With permission from Rockefeller University Press. B and C, from Q. Hu et al., Science 329:436–439, 2010. With permission from AAAS.)


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Cell_Biology_Alberts. A central challenge in cell biology is to understand how multiple individual molecular components collaborate to produce complex cell behaviors. The process of cell migration, which we describe in this final section, relies on the coordinated deployment of the components and processes that we have explored in this chapter: the dynamic assembly and disassembly of cytoskeletal polymers, the regulation and modification of their structure by polymer-associated proteins, and the actions of motor proteins moving along the polymers or exerting tension against them. How does the cell coordinate all these activities to define its polarity and enable it to crawl?


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Cell_Biology_Alberts. Many cells move by crawling over surfaces rather than by using cilia or flagella to swim. Predatory amoebae crawl continuously in search of food, and they can easily be observed to attack and devour smaller ciliates and flagellates in a drop of pond water (see Movie 1.4). In animals, almost all cell locomotion occurs by crawling, with the notable exception of swimming sperm. During embryogenesis, the structure of an animal is created by the migrations of individual cells to specific target locations and by the coordinated movements of whole epithelial sheets (discussed in Chapter 21). In vertebrates, neural crest cells are remarkable for their long-distance migrations from their site of origin in the neural tube to a variety of sites throughout the embryo (see Movie 21.5). Long-distance crawling is fundamental to the construction of the entire nervous system: it is in this way that the actin-rich growth cones at the advancing tips of developing axons travel to their eventual synaptic


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Cell_Biology_Alberts. is fundamental to the construction of the entire nervous system: it is in this way that the actin-rich growth cones at the advancing tips of developing axons travel to their eventual synaptic targets, guided by combinations of soluble signals and signals bound to cell surfaces and extracellular matrix along the way.


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Cell_Biology_Alberts. The adult animal also seethes with crawling cells. Macrophages and neutrophils crawl to sites of infection and engulf foreign invaders as a critical part of the innate immune response. Osteoclasts tunnel into bone, forming channels that are filled in by the osteoblasts that follow after them, in a continuous process of bone remodeling and renewal. Similarly, fibroblasts migrate through connective tissues, remodeling them where necessary and helping to rebuild damaged structures at sites of injury. In an ordered procession, the cells in the epithelial lining of the intestine travel up the sides of the intestinal villi, replacing absorptive cells lost at the tip of the villus. Unfortunately, cell crawling also has a role in many cancers, when cells in a primary tumor invade neighboring tissues and crawl into blood vessels or lymph vessels and then emerge at other sites in the body to form metastases.


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Cell_Biology_Alberts. Cell migration is a complex process that depends on the actin-rich cortex beneath the plasma membrane. Three distinct activities are involved: protrusion, in which the plasma membrane is pushed out at the front of the cell; attachment, in which the actin cytoskeleton connects across the plasma membrane to the substratum; and traction, in which the bulk of the trailing cytoplasm is drawn forward (Figure 16–75). In some crawling cells, such as keratocytes from the fish epidermis, these activities occur simultaneously, and the cells seem to glide forward smoothly without changing shape. In other cells, such as fibroblasts, these activities are more independent, and the locomotion is jerky and irregular.


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Cell_Biology_Alberts. The first step in locomotion, protrusion of a leading edge, frequently relies on forces generated by actin polymerization pushing the plasma membrane outward. Different cell types generate different types of protrusive structures, including filopodia (also known as microspikes) and lamellipodia. These are filled with dense cores of filamentous actin, which excludes membrane-enclosed organelles. The structures differ primarily in the way in which the actin is organized by actin-cross-linking proteins (see Figure 16–22). movement of unpolymerized actin


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Cell_Biology_Alberts. movement of unpolymerized actin Figure 16–75 A model of how forces generated in the actin-rich cortex move a cell forward. The actin-polymerizationdependent protrusion and firm attachment of a lamellipodium at the leading edge of the cell move the edge forward (green arrows at front) and stretch the actin cortex. Contraction at the rear of the cell propels the body of the cell forward (green arrow at back) to relax some of the tension (traction). New focal contacts are made at the front, and old ones are disassembled at the back as the cell crawls forward. The same cycle can be repeated, moving the cell forward in a stepwise fashion. Alternatively, all steps can be tightly coordinated, moving the cell forward smoothly. The newly polymerized cortical actin is shown in red.


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Cell_Biology_Alberts. Filopodia, formed by migrating growth cones of neurons and some types of fibroblasts, are essentially one-dimensional. They contain a core of long, bundled actin filaments, which are reminiscent of those in microvilli but longer and thinner, as well as more dynamic. Lamellipodia, formed by epithelial cells and fibroblasts, as well as by some neurons, are two-dimensional, sheetlike structures. They contain a cross-linked mesh of actin filaments, most of which lie in a plane parallel to the solid substratum. Invadopodia and related structures known as podosomes represent a third type of actin-rich protrusion. These extend in three dimensions and are important for cells to cross tissue barriers, as when a metastatic cancer cell invades the surrounding tissue. Invadopodia contain many of the same actin-regulatory components as filopodia and lamellipodia, and they also degrade the extracellular matrix, which requires the delivery of vesicles containing matrix-degrading proteases.


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Cell_Biology_Alberts. A distinct form of membrane protrusion called blebbing is often observed in vivo or when cells are cultured on a pliable extracellular matrix substratum. Blebs form when the plasma membrane detaches locally from the underlying actin cortex, thereby allowing cytoplasmic flow to push the membrane outward (Figure 16–76). Bleb formation also depends on hydrostatic pressure within the cell, which is generated by the contraction of actin and myosin assemblies. Once blebs have extended, actin filaments reassemble on the bleb membrane to form a new


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Cell_Biology_Alberts. Figure 16–76 Membrane bleb induced by disruption of the actin cortex. On the left is a light micrograph showing a spherical membrane protrusion or bleb induced by laser ablation of a small region of the actin cortex. The cortex is labeled green in the middle image by expression of GFP-actin. (Courtesy of Ewa Paluch.) actin cortex. Recruitment of myosin II and contraction of actin and myosin can then power retraction of membrane blebs. Alternatively, extension of new blebs from old ones can drive cell migration. Lamellipodia Contain All of the Machinery Required for Cell Motility


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Cell_Biology_Alberts. Lamellipodia Contain All of the Machinery Required for Cell Motility Lamellipodia have been particularly well studied in the epithelial cells of the epidermis of fish and frogs; these epithelial cells are known as keratocytes because of their abundant keratin filaments. These cells normally cover the animal by forming an epithelial sheet, and they are specialized to close wounds very rapidly, moving at rates of up to 30 μm/min. When cultured as individual cells, keratocytes assume a distinctive shape with a very large lamellipodium and a small, trailing cell body that is not attached to the substratum (Figure 16–77). Fragments of this lamellipodium can be sliced off with a micropipette. Although the fragments generally lack microtubules and membrane-enclosed organelles, they continue to crawl normally, looking like tiny keratocytes.


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Cell_Biology_Alberts. The dynamic behavior of actin filaments in keratocyte lamellipodia can be studied by labeling a small patch of actin and examining its fate. This reveals that, while the lamellipodia crawl forward, the actin filaments remain stationary with respect to the substratum. The actin filaments in the meshwork are mostly oriented with their plus ends facing forward. The minus ends are frequently attached to the sides of other actin filaments by Arp 2/3 complexes (see Figure 16–16), helping to form the two-dimensional web (Figure 16–78). The web as a whole is undergoing treadmilling, assembling at the front and disassembling at the back, reminiscent of the treadmilling that occurs in individual actin filaments discussed previously (see Figure 16–14).


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Cell_Biology_Alberts. Figure 16–78 Actin filament nucleation and web formation by the Arp 2/3 complex in lamellipodia. (A) A keratocyte with actin filaments labeled in red by fluorescent phalloidin and the Arp 2/3 complex labeled in green with an antibody against one of its subunits. The Arp 2/3 complex is highly concentrated near the front of the lamellipodium, where actin nucleation is most active. (B) Electron micrograph of a platinum-shadowed replica of the leading edge of a keratocyte, showing the dense actin filament meshwork. The labels denote areas enlarged in (C). (C) Close-up views of the marked regions of the actin web at the leading edge shown in (B). Numerous branched filaments can be seen, with the characteristic 70° angle formed when the Arp 2/3 complex nucleates a new actin filament off the side of a preexisting filament (see Figure 16–16). (From T. Svitkina and G. Borisy, J. Cell Biol. 145:1009–1026, 1999. With permission from the authors.)


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Cell_Biology_Alberts. J. Cell Biol. 145:1009–1026, 1999. With permission from the authors.) Figure 16–77 Migratory keratocytes from a fish epidermis. (A) Light micrographs of a keratocyte in culture, taken about 15 seconds apart. This cell is moving at about 15 μm/min (Movie 16.13 and see Movie 1.1). (B) Keratocyte seen by scanning electron microscopy, showing its broad, flat lamellipodium and small cell body, including the nucleus, carried up above the substratum at the rear. (C) Distribution of cytoskeletal filaments in this cell. Actin filaments (red) fill the large lamellipodium and are responsible for the cell’s rapid movement. Microtubules (green) and intermediate filaments (blue) are restricted to the regions close to the nucleus. (A and B, courtesy of Juliet Lee.) 954 Chapter 16: The Cytoskeleton