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Cell_Biology_Alberts. Maintenance of unidirectional motion by lamellipodia is thought to require the cooperation and mechanical integration of several factors. Filament nucleation is localized at the leading edge, with new actin filament growth occurring primarily in that location to push the plasma membrane forward. Most filament depolymerization occurs at sites located well behind the leading edge. Because cofilin (see Figure 16–20) binds cooperatively and preferentially to actin filaments containing ADP-actin (the D form), the new T-form filaments generated at the leading edge should be resistant to depolymerization by cofilin (Figure 16–79). As the filaments age and ATP hydrolysis proceeds, cofilin can efficiently disassemble the older filaments. Thus, the delayed ATP hydrolysis by filamentous actin is thought to provide the basis for a mechanism that maintains an efficient, unidirectional treadmilling process in the lamellipodium (Figure 16–80); it also explains the intracellular movement of bacterial


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Cell_Biology_Alberts. to provide the basis for a mechanism that maintains an efficient, unidirectional treadmilling process in the lamellipodium (Figure 16–80); it also explains the intracellular movement of bacterial pathogens such as Listeria (see Figure 16–25).


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Cell_Biology_Alberts. Myosin Contraction and Cell Adhesion Allow Cells to Pull Themselves Forward Forces generated by actin filament polymerization at the front of a migrating cell are transmitted to the underlying substratum to drive cell motion. For the leading Figure 16–79 Cofilin in lamellipodia. A keratocyte with actin filaments labeled in red by fluorescent phalloidin, and cofilin labeled in green with a fluorescent antibody. Although the dense actin meshwork reaches all the way through the lamellipodium, cofilin is not found at the very leading edge. (B) Close-up view of the region marked with the white rectangle in (A). The actin filaments closest to the leading edge, which are also the ones that have formed most recently and that are most likely to contain ATP-actin (rather than ADP-actin), are generally not associated with cofilin. (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. Figure 16–80 A model for protrusion of the actin meshwork at the leading edge. Two time points during advance of the lamellipodium are illustrated, with newly assembled structures at the later time point shown in a lighter color. Nucleation is mediated by the Arp 2/3 complex at the front. Newly nucleated actin filaments are attached to the sides of preexisting filaments, primarily at a 70° angle. Filaments elongate, pushing the plasma membrane forward because of some sort of anchorage of the array behind. At a steady rate, actin filament plus ends become capped. After newly polymerized actin subunits hydrolyze their bound ATP in the filament lattice, the filaments become susceptible to depolymerization by cofilin. This cycle causes a spatial separation between net filament assembly at the front and net filament disassembly at the rear, so that the actin filament network as a whole can move forward, even though the individual filaments within it remain stationary with respect to the


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Cell_Biology_Alberts. front and net filament disassembly at the rear, so that the actin filament network as a whole can move forward, even though the individual filaments within it remain stationary with respect to the substratum. Not all of the actin disassembles, however, and actin at the rear of the lamellipodium contributes to subsequent steps of migration together with myosin.


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Cell_Biology_Alberts. Figure 16–81 Contribution of myosin II to polarized cell motility.


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Cell_Biology_Alberts. (A) Myosin II bipolar filaments bind to actin filaments in the lamellipodial meshwork and cause network contraction. The myosin-driven reorientation of the actin filaments forms an actin bundle that recruits more myosin II and helps generate the contractile forces required for retraction of the trailing edge of the moving cell. (B) A fragment of the large lamellipodium of a keratocyte can be separated from the main cell body either by surgery with a micropipette or by treating the cell with certain drugs. Many of these fragments continue to move rapidly, with the same overall cytoskeletal organization as the intact keratocytes. Actin (blue) forms a protrusive meshwork at the front of the fragment. Myosin II (pink) is gathered into a band at the rear. (From A. Verkhovsky et al., Curr. Biol. 9:11–20, 1999. With permission from Elsevier.) edge of a migrating cell to advance, protrusion of the membrane must be followed by adhesion to the substratum at the front. Conversely, in order for


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Cell_Biology_Alberts. 9:11–20, 1999. With permission from Elsevier.) edge of a migrating cell to advance, protrusion of the membrane must be followed by adhesion to the substratum at the front. Conversely, in order for the cell body to follow, contraction must be coupled with de-adhesion at the rear of the cell. The processes contributing to migration are therefore tightly regulated in space and time, with actin polymerization, dynamic adhesions, and myosin contraction being employed to coordinate movement. Myosin II operates in at least two ways to assist cell migration. The first is by helping to connect the actin cytoskeleton to the substratum through integrin-mediated adhesions. Forces generated by both actin polymerization and myosin activity create tension at attachment sites, promoting their maturation into focal adhesions, which are dynamic assemblies of structural and signaling proteins that link the migrating cell to the extracellular matrix (see Figure 19–59). A second mechanism involves bipolar


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Cell_Biology_Alberts. focal adhesions, which are dynamic assemblies of structural and signaling proteins that link the migrating cell to the extracellular matrix (see Figure 19–59). A second mechanism involves bipolar myosin II filaments, which associate with the actin filaments at the rear of the lamellipodium and pull them into a new orientation—from nearly perpendicular to the leading edge to almost parallel to the leading edge. This sarcomere-like contraction prevents protrusion, and it pinches in the sides of the locomoting lamellipodium, helping to gather in the sides of the cell as it moves forward (Figure 16–81).


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Cell_Biology_Alberts. Actin-mediated protrusions can only push the leading edge of the cell forward if there are strong interactions between the actin network and the focal adhesions that link the cell to the substrate. When these interactions are disengaged, polymerization pressure at the leading edge and myosin-dependent contraction cause the actin network to slip back, resulting in a phenomenon known as retrograde flow (Figure 16–82).


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Cell_Biology_Alberts. The traction forces generated by locomoting cells exert a significant pull on the substratum. By growing cells on a surface coated with tiny flexible posts, the force exerted on the substratum can be calculated by measuring the deflection of each post from its vertical position (Figure 16–83). In a living animal, most crawling cells move across a semiflexible substratum made of extracellular matrix, which can be deformed and rearranged by these cell forces. Conversely, mechanical tension or stretching applied externally to a cell will cause it to assemble stress fibers and focal adhesions, and become more contractile. Although poorly understood, this two-way mechanical interaction between cells and their physical environment is thought to help vertebrate tissues organize themselves. Cell Polarization Is Controlled by Members of the Rho Protein Family


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Cell_Biology_Alberts. Cell Polarization Is Controlled by Members of the Rho Protein Family Cell migration requires long-distance communication and coordination between one end of a cell and the other. During directed migration, it is important that the front end of the cell remain structurally and functionally distinct from the back end. In addition to driving local mechanical processes such as protrusion at the front and retraction at the rear, the cytoskeleton is responsible for coordinating cell shape, organization, and mechanical properties from one end of the cell to the other, a distance that is typically tens of micrometers for animal cells.


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Cell_Biology_Alberts. In many cases, including but not limited to cell migration, large-scale cytoskeletal coordination takes the form of the establishment of cell polarity, where a cell builds different structures with distinct molecular components at the front versus the back, or at the top versus the bottom. Cell locomotion requires an initial polarization of the cell to set it off in a particular direction. Carefully controlled cell-polarization processes are also required for oriented cell divisions in tissues and for formation of a coherent, organized multicellular structure. Genetic studies in yeast, flies, and worms have provided most of our current understanding of the molecular basis of cell polarity. The mechanisms that generate cell polarity in vertebrates are only beginning to be explored. In all known cases, however, the cytoskeleton has a central role, and many of the molecular components have been evolutionarily conserved.


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Cell_Biology_Alberts. The establishment of many kinds of cell polarity depends on the local regulation of the actin cytoskeleton by external signals. Many of these signals seem to converge inside the cell on a group of closely related monomeric GTPases that are members of the Rho protein family—Cdc42, Rac, and Rho. Like other monomeric GTPases, the Rho proteins act as molecular switches that cycle between an active GTP-bound state and an inactive GDP-bound state (see Figure 3–66). Activation of Cdc42 on the inner surface of the plasma membrane triggers actin polymerization and bundling to form filopodia. Activation of Rac promotes actin polymerization at the cell periphery, leading to the formation of sheetlike lamellipodial extensions. Activation of Rho promotes both the bundling of actin filaments with myosin II filaments into stress fibers and the clustering of integrins


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Cell_Biology_Alberts. Figure 16–82 Control of cell– substratum adhesion at the leading edge of a migrating cell. (A) Actin monomers assemble on the barbed end of actin filaments at the leading edge. Transmembrane integrin proteins (blue) help form focal adhesions that link the cell membrane to the substrate. (B) If there is no interaction between the actin filaments and focal adhesions, the actin filament is driven rearward by newly assembled actin. Myosin motors (green) also contribute to filament movement. (C) Interactions between actin-binding adaptor proteins (brown) and integrins link the actin cytoskeleton to the substratum. Myosin-mediated contractile forces are then transmitted through the focal adhesion to generate traction on the extracellular matrix, and new actin polymerization drives the leading edge forward in a protrusion.


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Cell_Biology_Alberts. Figure 16–83 Traction forces exerted by a motile cell. (A) Tiny flexible pillars attached to the substratum bend in response to traction forces. (B) Scanning electron micrograph of a cell on a substratum coated with pillars that are 6.1 μm in height. Pillar deflections are used to calculate force vectors corresponding to inward pulling forces on the underlying substratum. (Adapted from J. Fu et al., Nat. Methods 7:733–736, 2010. With permission from Macmillan Publishers.) and associated proteins to form focal adhesions (Figure 16–84). These dramatic and complex structural changes occur because each of these three molecular switches has numerous downstream target proteins that affect actin organization and dynamics.


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Cell_Biology_Alberts. Some key targets of activated Cdc42 are members of the WASp protein family. Human patients deficient in WASp suffer from Wiskott-Aldrich Syndrome, a severe form of immunodeficiency in which immune system cells have abnormal actin-based motility and platelets do not form normally. Although WASp itself is expressed only in blood cells and immune system cells, other more ubiquitous versions enable activated Cdc42 to enhance actin polymerization in many cell types. WASp proteins can exist in an inactive folded conformation and an activated open conformation. Association with Cdc42-GTP stabilizes the open form of WASp, enabling it to bind to the Arp 2/3 complex and strongly enhance its actin-nucleating activity (see Figure 16–16). In this way, activation of Cdc42 increases actin nucleation.


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Cell_Biology_Alberts. Rac-GTP also activates WASp family members. Additionally, it activates the cross-linking activity of the gel-forming protein filamin and inhibits the contractile activity of the motor protein myosin II. It thereby stabilizes lamellipodia and inhibits the formation of contractile stress fibers (Figure 16–85A). Rho-GTP has a very different set of targets. Instead of activating the Arp 2/3 complex to build actin networks, Rho-GTP turns on formin proteins to construct parallel actin bundles. At the same time, Rho-GTP activates a protein kinase that indirectly inhibits the activity of cofilin, leading to actin filament stabilization. The same protein kinase inhibits a phosphatase acting on myosin light chains (see Figure 16–39). The consequent increase in the net amount of myosin light chain phosphorylation increases the amount of contractile myosin motor protein activity in the cell, enhancing the formation of tension-dependent structures such as stress fibers (Figure 16–85B).


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Cell_Biology_Alberts. In some cell types, Rac-GTP activates Rho, usually at a rate that is slow compared to Rac’s activation of the Arp 2/3 complex. This enables cells to use the Rac pathway to build a new actin structure while subsequently activating the Rho pathway to generate a contractility that builds up tension in this structure. This occurs, for example, during the formation and maturation of cell–cell contacts.


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Cell_Biology_Alberts. Figure 16–84 The dramatic effects of Cdc42, Rac, and Rho on actin organization in fibroblasts. In each case, the actin filaments have been labeled with fluorescent phalloidin. (A) Serum-starved fibroblasts have actin filaments primarily in the cortex, and relatively few stress fibers. (B) Microinjection of a constitutively activated form of Cdc42 causes the protrusion of many long filopodia at the cell periphery. (C) Microinjection of a constitutively activated form of Rac, a closely related monomeric GTPase, causes the formation of an enormous lamellipodium that extends from the entire circumference of the cell. (D) Microinjection of a constitutively activated form of Rho causes the rapid assembly of many prominent stress fibers. (From A. Hall, Science 279:509–514, 1998. With permission from AAAS.)


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Cell_Biology_Alberts. (D) Microinjection of a constitutively activated form of Rho causes the rapid assembly of many prominent stress fibers. (From A. Hall, Science 279:509–514, 1998. With permission from AAAS.) As we will explore in more detail below, the communication between the Rac and Rho pathways also facilitates maintenance of the large-scale differences between the cell front and the cell rear during migration. Extracellular Signals Can Activate the Three Rho Protein Family Members


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Cell_Biology_Alberts. The activation of the monomeric GTPases Rho, Rac, and Cdc42 occurs through an exchange of GTP for a tightly bound GDP molecule, catalyzed by guanine nucleotide exchange factors (GEFs). Of the many GEFs that have been identified in the human genome, some are specific for an individual Rho family GTPase, whereas others seem to act on multiple family members. Different GEFs are restricted to specific tissues and even specific subcellular locations, and they are sensitive to distinct kinds of regulatory inputs. GEFs can be activated by extracellular cues through cell-surface receptors, or in response to intracellular signals. GEFs may also act as scaffolds that direct GTPases to downstream effectors. Interestingly, several of the Rho family GEFs associate with the growing ends of microtubules by binding to one of the +TIPs. This provides a connection between the dynamics of the microtubule cytoskeleton and the large-scale organization of the actin cytoskeleton; such a connection is


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Cell_Biology_Alberts. by binding to one of the +TIPs. This provides a connection between the dynamics of the microtubule cytoskeleton and the large-scale organization of the actin cytoskeleton; such a connection is important for the overall integration of cell shape and movement.


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Cell_Biology_Alberts. External Signals Can Dictate the Direction of Cell Migration Chemotaxis is the movement of a cell toward or away from a source of some diffusible chemical. These external signals act through Rho family proteins to set up large-scale cell polarity by influencing the organization of the cell motility apparatus. One well-studied example is the chemotactic movement of a class of white blood cells, called neutrophils, toward a source of bacterial infection. Receptor proteins on the surface of neutrophils enable them to detect very low concentrations of N-formylated peptides that are derived from bacterial proteins (only prokaryotes begin protein synthesis with N-formylmethionine). Using these receptors, neutrophils are guided to bacterial targets by their ability to detect a difference of only 1% in the concentration of these diffusible peptides on one side of the cell versus the other (Figure 16–86A).


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Cell_Biology_Alberts. In this case, and in the chemotaxis of Dictyostelium amoebae toward a source of cyclic AMP, binding of the chemoattractant to its G-protein-coupled receptor activates phosphoinositide 3-kinases (PI3Ks) (see Figure 15–52), which generate a signaling molecule [PI(3,4,5)P3] that in turn activates the Rac GTPase. Rac


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Cell_Biology_Alberts. Figure 16–85 The contrasting effects of Rac and Rho activation on actin organization. (A) Activation of the small GTPase Rac leads to alterations in actin accessory proteins that tend to favor the formation of actin networks, as in lamellipodia. Several different pathways contribute independently. Rac-GTP activates members of the WASp protein family, which in turn activate actin nucleation and branched web formation by the Arp 2/3 complex. In a parallel pathway, Rac-GTP activates a protein kinase, PAK, which has several targets including the web-forming cross-linker filamin, which is activated by phosphorylation, and the myosin light chain kinase (MLCK), which is inhibited by phosphorylation. Inhibition of MLCK results in decreased phosphorylation of the myosin regulatory light chain and leads to myosin II filament disassembly and a decrease in contractile activity. In some cells, PAK also directly inhibits myosin II activity by phosphorylation of the myosin heavy chain (MHC). (B)


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Cell_Biology_Alberts. leads to myosin II filament disassembly and a decrease in contractile activity. In some cells, PAK also directly inhibits myosin II activity by phosphorylation of the myosin heavy chain (MHC). (B) Activation of the related GTPase Rho leads to nucleation of actin filaments by formins and increases contraction by myosin II, promoting the formation of contractile actin bundles such as stress fibers. Activation of myosin II by Rho requires a Rho-dependent protein kinase called Rock. This kinase inhibits the phosphatase that removes the activating phosphate groups from myosin II light chains (MLC); it may also directly phosphorylate the myosin light chains in some cell types. Rock also activates other protein kinases, such as LIM kinase, which in turn contributes to the formation of stable contractile actin filament bundles by inhibiting the actin depolymerizing factor cofilin. A similar signaling pathway is important for forming the contractile ring necessary for cytokinesis (see Figure


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Cell_Biology_Alberts. actin filament bundles by inhibiting the actin depolymerizing factor cofilin. A similar signaling pathway is important for forming the contractile ring necessary for cytokinesis (see Figure 17–44).


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Cell_Biology_Alberts. 5 µm neutrophil Gi G12/13 Rho PI(3,4,5)P3 Rac Rac dominates, polymerization (protrusion) Rho dominates, actin–myosin contraction chemoattractant receptor bacterium back front (A) (B) then activates the Arp 2/3 complex leading to lamellipodial protrusion. Through an unknown mechanism, accumulation of the polarized actin web at the leading edge causes further local enhancement of PI3K activity in a positive feedback loop, strengthening the induction of protrusion. The PI(3,4,5)P3 that activates Rac cannot diffuse far from its site of synthesis, since it is rapidly converted back into PI(4,5)P2 by a constitutively active lipid phosphatase. At the same time, binding of the chemoattractant ligand to its receptor activates another signaling pathway that turns on Rho and enhances myosin-based contractility. The two processes directly inhibit each other, such that Rac activation dominates in the front of the cell and Rho activation dominates in the rear (Figure 16–86B). This enables the cell


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Cell_Biology_Alberts. The two processes directly inhibit each other, such that Rac activation dominates in the front of the cell and Rho activation dominates in the rear (Figure 16–86B). This enables the cell to maintain its functional polarity with protrusion at the leading edge and contraction at the back.


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Cell_Biology_Alberts. Nondiffusible chemical cues attached to the extracellular matrix or to the surface of cells can also influence the direction of cell migration. When these signals activate receptors, they can cause increased cell adhesion and directed actin polymerization. Most long-distance cell migrations in animals, including neuralcrest-cell migration and the travels of neuronal growth cones, depend on a combination of diffusible and nondiffusible signals to steer the locomoting cells or growth cones to their proper destinations.


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Cell_Biology_Alberts. The interconnected cytoskeleton is crucial for cell migration. Although movement is driven primarily by actin polymerization and myosin contractility, septins and intermediate filaments also participate. For example, vimentin intermediate filament networks associate with integrins at focal adhesions, and vimentin-deficient fibroblasts display impaired mechanical stability, migration, and contractile capacity. Furthermore, disruption of linker proteins that connect different cytoskeletal elements, including several plakins and KASH proteins, leads to defects in cell polarization and migration. Thus, interactions among cytoplasmic filament systems, as well as mechanical linkage to the nucleus, are required for complex, whole-cell behaviors such as migration.


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Cell_Biology_Alberts. Cells also use microtubules to help organize persistent movement in a specific direction. In many locomoting cells, the position of the centrosome is influenced by the location of protrusive actin polymerization. Activation of receptors on the protruding front edge of a cell might locally activate dynein motor proteins that move the centrosome by pulling on its microtubules. Several effector proteins downstream of Rac and Rho modulate microtubule dynamics directly: for example, a protein kinase activated by Rac can phosphorylate (and thereby inhibit) the tubulin-binding protein stathmin (see Panel 16–4), thereby stabilizing microtubules.


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Cell_Biology_Alberts. Figure 16–86 Neutrophil polarization and chemotaxis. (A) The pipette tip at the right is leaking a small amount of the bacterial peptide formyl-Met-Leu-Phe, which is recognized by the human neutrophil as the product of a foreign invader. The neutrophil quickly extends a new lamellipodium toward the source of the chemoattractant peptide (top). It then extends this lamellipodium and polarizes its cytoskeleton so that contractile myosin II is located primarily at the rear, opposite the position of the lamellipodium (middle). Finally, the cell crawls toward the source of the peptide (bottom). If a real bacterium were the source of the peptide, rather than an investigator’s pipette, the neutrophil would engulf the bacterium and destroy it (see also Figure 16–3 and Movie 16.14). (B) Binding of bacterial molecules to G-protein-coupled receptors on the neutrophil stimulates directed motility. These receptors are found all over the surface of the cell, but are more likely to be bound to the


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Cell_Biology_Alberts. bacterial molecules to G-protein-coupled receptors on the neutrophil stimulates directed motility. These receptors are found all over the surface of the cell, but are more likely to be bound to the bacterial ligand at the front. Two distinct signaling pathways contribute to the cell’s polarization. At the front of the cell, stimulation of the Rac pathway leads, via the trimeric G protein Gi, to growth of protrusive actin networks. Second messengers within this pathway are short-lived, so protrusion is limited to the region of the cell closest to the stimulant. The same receptor also stimulates a second signaling pathway, via the trimeric G proteins G12 and G13, that triggers the activation of Rho. The two pathways are mutually antagonistic. Since Rac-based protrusion is active at the front of the cell, Rho is activated only at the rear of the cell, stimulating contraction of the cell rear and assisting directed movement. (A, from


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Cell_Biology_Alberts. O.D. Weiner et al., Nat. Cell Biol. 1:75–81, 1999. With permission from Macmillan Publishers Ltd.) In turn, microtubules influence actin rearrangements and cell adhesion. The centrosome nucleates a large number of dynamic microtubules, and its repositioning means that the plus ends of many of these microtubules extend into the protrusive region of the cell. Direct interactions with microtubules help guide focal adhesion dynamics in migrating cells. Microtubules might also influence actin filament formation by delivering Rac-GEFs that bind to the +TIPs traveling on growing microtubule ends. Microtubules also transport cargoes to and from the focal adhesions, thereby affecting their signaling and disassembly. Thus, microtubules reinforce the polarity information that the actin cytoskeleton receives from the outside world, allowing a sensitive response to weak signals and enabling motility to persist in the same direction for a prolonged period.


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Cell_Biology_Alberts. Whole-cell movements and the large-scale shaping and structuring of cells require the coordinated activities of all three basic filament systems along with a large variety of cytoskeletal accessory proteins, including motor proteins. Cell crawling—a widespread behavior important in embryonic development and also in wound healing, tissue maintenance, and immune system function in the adult animal— is a prime example of such complex, coordinated cytoskeletal action. For a cell to crawl, it must generate and maintain an overall structural polarity, which is influenced by external cues. In addition, the cell must coordinate protrusion at the leading edge (by assembly of new actin filaments), adhesion of the newly protruded part of the cell to the substratum, and forces generated by molecular motors to bring the cell body forward. Which statements are true? Explain why or why not.


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Cell_Biology_Alberts. Which statements are true? Explain why or why not. 16–1 The role of ATP hydrolysis in actin polymerization is similar to the role of GTP hydrolysis in tubulin polymerization: both serve to weaken the bonds in the polymer and thereby promote depolymerization. 16–2 Motor neurons trigger action potentials in muscle cell membranes that open voltage-sensitive Ca2+ channels in T tubules, allowing extracellular Ca2+ to enter the cytosol, bind to troponin C, and initiate rapid muscle contraction. 16–3 In most animal cells, minus-end directed microtubule motors deliver their cargo to the periphery of the cell, whereas plus-end directed microtubule motors deliver their cargo to the interior of the cell. Discuss the following problems.


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Cell_Biology_Alberts. Discuss the following problems. 16–4 The concentration of actin in cells is 50–100 times greater than the critical concentration observed for pure actin in a test tube. How is this possible? What prevents the actin subunits in cells from polymerizing into filaments? Why is it advantageous to the cell to maintain such a large pool of actin subunits? 16–5 Detailed measurements of sarcomere length and tension during isometric contraction in striated muscle provided crucial early support for the sliding-filament 2.0 2.2 1.6 1.3 I 3.6 How is the cell cortex regulated locally and globally to coordinate its activities at different places on the cell surface? What determines, for example, where filopodia form? How are actin-regulatory proteins controlled spatially in the cytoplasm to generate multiple distinct types of actin arrays in the same cell? Are there biologically important processes occurring inside a microtubule?


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Cell_Biology_Alberts. Are there biologically important processes occurring inside a microtubule? How can we account for the fact that there are many different kinesins and myosins in the cytoplasm but only one dynein? Mutations in the nuclear lamin proteins cause a large number of diseases called laminopathies. What do we not understand about the nuclear lamina that could account for this fact? Figure Q16–1 Tension as a function of sarcomere length during isometric contraction (Problem 16–5).


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Cell_Biology_Alberts. Figure Q16–1 Tension as a function of sarcomere length during isometric contraction (Problem 16–5). model of muscle contraction. Based on your understanding of the sliding-filament model and the structure of a sarcomere, propose a molecular explanation for the relationship of tension to sarcomere length in the portions of Figure Q16–1 marked I, II, III, and IV. (In this muscle, the length of the myosin filament is 1.6 μm, and the lengths of the actin thin filaments that project from the Z discs are 1.0 μm.) 16–6 At 1.4 mg/mL pure tubulin, microtubules grow at a rate of about 2 μm/min. At this growth rate, how many αβ-tubulin dimers (8 nm in length) are added to the ends of a microtubule each second? Figure Q16–2 Model for microtubule nucleation by pure αβ-tubulin dimers (Problem 16–7).


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Cell_Biology_Alberts. Figure Q16–2 Model for microtubule nucleation by pure αβ-tubulin dimers (Problem 16–7). 16–7 A solution of pure αβ-tubulin dimers is thought to nucleate microtubules by forming a linear protofilament about seven dimers in length. At that point, the probabilities that the next αβ-dimer will bind laterally or to the end of the protofilament are about equal. The critical event for microtubule formation is thought to be the first lateral association (Figure Q16–2). How does lateral association promote the subsequent rapid formation of a microtubule? 16–8 How does a centrosome “know” when it has found the center of the cell?


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Cell_Biology_Alberts. 16–8 How does a centrosome “know” when it has found the center of the cell? 16–9 The movements of single motor-protein molecules can be analyzed directly. Using polarized laser light, it is possible to create interference patterns that exert a centrally directed force, ranging from zero at the center to a few piconewtons at the periphery (about 200 nm from the center). Individual molecules that enter the interference pattern are rapidly pushed to the center, allowing them to be captured and moved at the experimenter’s discretion.


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Cell_Biology_Alberts. Using such “optical tweezers,” single kinesin molecules can be positioned on a microtubule that is fixed to a coverslip. Although a single kinesin molecule cannot be seen optically, it can be tagged with a silica bead and tracked indirectly by following the bead (Figure Q16–3A). In the absence of ATP, the kinesin molecule remains at the center of the interference pattern, but with ATP it moves toward the plus end of the microtubule. As kinesin moves along the microtubule, it encounters the force of the interference pattern, which simulates the load kinesin carries during its actual function in the cell. Moreover, the pressure against the silica bead counters the effects of Brownian (thermal) motion, so that the position of the bead more accurately reflects the position of the kinesin molecule on the microtubule. A trace of the movements of a kinesin molecule along a microtubule is shown in Figure Q16–3B.


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Cell_Biology_Alberts. A trace of the movements of a kinesin molecule along a microtubule is shown in Figure Q16–3B. A. As shown in Figure Q16–3B, all movement of kine-sin is in one direction (toward the plus end of the micro-tubule). What supplies the free energy needed to ensure a unidirectional movement along the microtubule? b. What is the average rate of movement of kinesin along the microtubule? C. What is the length of each step that a kinesin takes as it moves along a microtubule? D. From other studies it is known that kinesin has two globular domains that can each bind to β-tubulin, and that kinesin moves along a single protofilament in a microtubule. In each protofilament, the β-tubulin subunit repeats at 8-nm intervals. Given the step length and the interval between β-tubulin subunits, how do you suppose a kine-sin molecule moves along a microtubule? E. Is there anything in the data in Figure Q16–3B that tells you how many ATP molecules are hydrolyzed per step?


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Cell_Biology_Alberts. E. Is there anything in the data in Figure Q16–3B that tells you how many ATP molecules are hydrolyzed per step? 16–10 A mitochondrion 1 μm long can travel the 1 meter length of the axon from the spinal cord to the big toe in a day. The Olympic men’s freestyle swimming record for 200 meters is 1.75 minutes. In terms of body lengths per day, who is moving faster: the mitochondrion or the Olympic record holder? (Assume that the swimmer is 2 meters tall.) 16–11 Cofilin preferentially binds to older actin filaments and promotes their disassembly. How does cofilin distinguish old filaments from new ones? 16–12 Why is it that intermediate filaments have identical ends and lack polarity, whereas actin filaments and microtubules have two distinct ends with a defined polarity? 16–13 How is the unidirectional motion of a lamellipodium maintained?


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Cell_Biology_Alberts. 16–13 How is the unidirectional motion of a lamellipodium maintained? Figure Q16–3 Movement of kinesin along a microtubule (Problem 16–9). (A) Experimental set-up, with kinesin linked to a silica bead, moving along a microtubule. (B) Position of kinesin (as visualized by the position of the silica bead) relative to the center of the interference pattern, as a function of time of movement along the microtubule. The jagged nature of the trace results from Brownian motion of the bead. Bray D (2001) Cell Movements: From Molecules to Motility, 2nd ed. New York: Garland Science. Howard J (2001) Mechanics of Motor Proteins and the Cytoskeleton. Sunderland, MA: Sinauer. Kavallaris M (2012) Cytoskeleton and Human Disease. New York: Springer, Humana Press. Function and Origin of the Cytoskeleton Garner EC, Bernard R, Wang W et al. (2011) Coupled, circumferential motions of the cell wall synthesis machinery and MreB filaments in B. subtilis. Science 333, 222–225.


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Cell_Biology_Alberts. Garner EC, Bernard R, Wang W et al. (2011) Coupled, circumferential motions of the cell wall synthesis machinery and MreB filaments in B. subtilis. Science 333, 222–225. Garner EC, Campbell CS & Mullins RD (2004) Dynamic instability in a DNA-segregating prokaryotic actin homolog. Science 306, 1021–1025. Hill TL & Kirschner MW (1982) Bioenergetics and kinetics of microtubule and actin filament assembly-disassembly. Int. Rev. Cytol. 78, 1–125. Jones LJ, Carballido-López R & Errington J (2001) Control of cell shape in bacteria: helical, actin-like filaments in Bacillus subtilis. Cell 104, 913–922. Luby-Phelps K (2000) Cytoarchitecture and physical properties of cytoplasm: volume, viscosity, diffusion, intracellular surface area. Int. Rev. Cytol. 192, 189–221. Oosawa F & Asakura S (1975) Thermodynamics of the Polymerization of Protein, pp. 41–55 and 90–108. New York: Academic Press.


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Cell_Biology_Alberts. Oosawa F & Asakura S (1975) Thermodynamics of the Polymerization of Protein, pp. 41–55 and 90–108. New York: Academic Press. Osawa M, Anderson DE & Erickson HP (2009) Curved FtsZ protofilaments generate bending forces on liposome membranes. EMBO J. 28, 3476–3484. Pauling L (1953) Aggregation of globular proteins. Discuss. Faraday Soc. 13, 170–176. Purcell EM (1977) Life at low Reynolds number. Am. J. Phys. 45, 3–11. Theriot JA (2013) Why are bacteria different from eukaryotes? BMC Biol. 11, 119. Fehon RG, McClatchey AI & Bretscher A (2010) Organizing the cell cortex: the role of ERM proteins. Nat. Rev. Mol. Cell Biol. 11, 276–287. Mullins RD, Heuser JA & Pollard TD (1998) The interaction of Arp2/3 complex with actin: nucleation, high affinity pointed end capping, and formation of branching networks of filaments. Proc. Natl Acad. Sci. U.S.A. 95, 6181–6186. Zigmond SH (2004) Formin-induced nucleation of actin filaments. Curr. Opin. Cell Biol. 16, 99–105.


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Cell_Biology_Alberts. Zigmond SH (2004) Formin-induced nucleation of actin filaments. Curr. Opin. Cell Biol. 16, 99–105. Cooke R (2004) The sliding filament model: 1972–2004. J. Gen. Physiol. 123, 643–656. Hammer JA 3rd & Sellers JR (2011) Walking to work: roles for class V myosins as cargo transporters. Nat. Rev. Mol. Cell Biol. 13, 13–26. Howard J (1997) Molecular motors: structural adaptations to cellular functions. Nature 389, 561–567. Rice S, Lin AW, Safer D et al. (1999) A structural change in the kinesin motor protein that drives motility. Nature 402, 778–784. Vikstrom KL & Leinwand LA (1996) Contractile protein mutations and heart disease. Curr. Opin. Cell Biol. 8, 97–105. Wells AL, Lin AW, Chen LQ et al. (1999) Myosin VI is an actin-based motor that moves backwards. Nature 401, 505–508. Yildiz A, Forkey JN, McKinney SA et al. (2003) Myosin V walks hand-over-hand: single fluorophore imaging with 1.5-nm localization. Science 300, 2061–2065.


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Cell_Biology_Alberts. Yildiz A, Forkey JN, McKinney SA et al. (2003) Myosin V walks hand-over-hand: single fluorophore imaging with 1.5-nm localization. Science 300, 2061–2065. Aldaz H, Rice LM, Stearns T & Agard DA (2005) Insights into microtubule nucleation from the crystal structure of human gammatubulin. Nature 435, 523–527. Brouhard GJ, Stear JH, Noetzel TL et al. (2008) XMAP215 is a processive microtubule polymerase. Cell 132, 79–88. Dogterom M & Yurke B (1997) Measurement of the force-velocity relation for growing microtubules. Science 278, 856–860. Doxsey S, McCollum D & Theurkauf W (2005) Centrosomes in cellular regulation. Annu. Rev. Cell Dev. Biol. 21, 411–434. Galjart N (2010) Plus-end-tracking proteins and their interactions at microtubule ends. Curr. Biol. 20, R528–R537. Hotani H & Horio T (1988) Dynamics of microtubules visualized by darkfield microscopy: treadmilling and dynamic instability. Cell Motil. Cytoskeleton 10, 229–236.


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Cell_Biology_Alberts. Hotani H & Horio T (1988) Dynamics of microtubules visualized by darkfield microscopy: treadmilling and dynamic instability. Cell Motil. Cytoskeleton 10, 229–236. Howard J, Hudspeth AJ & Vale RD (1989) Movement of microtubules by single kinesin molecules. Nature 342, 154–158. Kerssemakers JW, Munteanu EL, Laan L et al. (2006) Assembly dynamics of microtubules at molecular resolution. Nature 442, 709–712. Kikkawa M (2013) Big steps toward understanding dynein. J. Cell Biol. 202, 15–23. Mitchison T & Kirschner M (1984) Dynamic instability of microtubule growth. Nature 312, 237–242. Reck-Peterson SL, Yildiz A, Carter AP et al. (2006) Single-molecule analysis of dynein processivity and stepping behavior. Cell 126, 335–348. Sharp DJ & Ross JL (2012) Microtubule-severing enzymes at the cutting edge. J. Cell Sci. 125, 2561–2569. Singla V & Reiter JF (2006) The primary cilium as the cell’s antenna: signaling at a sensory organelle. Science 313, 629–633.


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Cell_Biology_Alberts. Singla V & Reiter JF (2006) The primary cilium as the cell’s antenna: signaling at a sensory organelle. Science 313, 629–633. Stearns T & Kirschner M (1994) In vitro reconstitution of centrosome assembly and function: the central role of gamma-tubulin. Cell 76, 623–637. Svoboda K, Schmidt CF, Schnapp BJ & Block SM (1993) Direct observation of kinesin stepping by optical trapping interferometry. Nature 365, 721–727. Verhey KJ, Kaul N & Soppina V (2011) Kinesin assembly and movement in cells. Annu. Rev. Biophys. 40, 267–288. Helfand BT, Chang L & Goldman RD (2003) The dynamic and motile properties of intermediate filaments. Annu. Rev. Cell Dev. Biol. 19, 445–467. Isermann P & Lammerding J (2013) Nuclear mechanics and mechanotransduction in health and disease. Curr. Biol. 23, R1113–R1121. Saarikangas J & Barral Y (2011) The emerging functions of septins in metazoans. EMBO Rep. 12, 1118–1126.


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Cell_Biology_Alberts. Saarikangas J & Barral Y (2011) The emerging functions of septins in metazoans. EMBO Rep. 12, 1118–1126. Abercrombie M (1980) The crawling movement of metazoan cells. Proc. R. Soc. Lond. B 207, 129–147. Gardel ML, Schneider IC, Aratyn-Schaus Y & Waterman CM (2010) Mechanical integration of actin and adhesion dynamics in cell migration. Annu. Rev. Cell Dev. Biol. 26, 315–333. Lo CM, Wang HB, Dembo M & Wang YL (2000) Cell movement is guided by the rigidity of the substrate. Biophys. J. 79, 144–152. Madden K & Snyder M (1998) Cell polarity and morphogenesis in budding yeast. Annu. Rev. Microbiol. 52, 687–744. Parent CA & Devreotes PN (1999) A cell’s sense of direction. Science 284, 765–770. Pollard TD & Borisy GG (2003) Cellular motility driven by assembly and disassembly of actin filaments. Cell 112, 453–465. Rafelski SM & Theriot JA (2004) Crawling toward a unified model of cell mobility: spatial and temporal regulation of actin dynamics. Annu. Rev. Biochem. 73, 209–239.


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Cell_Biology_Alberts. Rafelski SM & Theriot JA (2004) Crawling toward a unified model of cell mobility: spatial and temporal regulation of actin dynamics. Annu. Rev. Biochem. 73, 209–239. Ridley A (2011) Life at the leading edge. Cell 145, 1012–1022. Vitriol EA & Zheng JQ (2012) Growth cone travel in space and time: the cellular ensemble of cytoskeleton, adhesion, and membrane. Neuron 73, 1068–1081. Weiner OD (2002) Regulation of cell polarity during eukaryotic chemotaxis: the chemotactic compass. Curr. Opin. Cell Biol. 14, 196–202. The Cell Cycle 17 The only way to make a new cell is to duplicate a cell that already exists. This simple fact, first established in the middle of the nineteenth century, carries with it a profound message for the continuity of life. All living organisms, from the unicellular bacterium to the multicellular mammal, are products of repeated rounds of cell growth and division extending back in time to the beginnings of life on Earth over three billion years ago.


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Cell_Biology_Alberts. A cell reproduces by performing an orderly sequence of events in which it duplicates its contents and then divides in two. This cycle of duplication and division, known as the cell cycle, is the essential mechanism by which all living things reproduce. In unicellular species, such as bacteria and yeasts, each cell division produces a complete new organism. In multicellular species, long and complex sequences of cell divisions are required to produce a functioning organism. Even in the adult body, cell division is usually needed to replace cells that die. In fact, each of us must manufacture many millions of cells every second simply to survive: if all cell division were stopped—by exposure to a very large dose of x-rays, for example—we would die within a few days.


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Cell_Biology_Alberts. The details of the cell cycle vary from organism to organism and at different times in an organism’s life. Certain characteristics, however, are universal. At a minimum, the cell must accomplish its most fundamental task: the passing on of its genetic information to the next generation of cells. To produce two genetically identical daughter cells, the DNA in each chromosome must first be faithfully replicated to produce two complete copies. The replicated chromosomes must then be accurately distributed (segregated) to the two daughter cells, so that each receives a copy of the entire genome (Figure 17–1). In addition to duplicating their genome, most cells also duplicate their other organelles and macromolecules; otherwise, daughter cells would get smaller with each division. To maintain their size, dividing cells must coordinate their growth (that is, their increase in cell mass) with their division.


Cell_Biology_Alberts

Cell_Biology_Alberts. This chapter describes the events of the cell cycle and how they are controlled and coordinated. We begin with a brief overview of the cell cycle. We then describe the cell-cycle control system, a complex network of regulatory proteins that triggers the different events of the cycle. We next consider in detail the major stages of the cell cycle, in which the chromosomes are duplicated and then segregated into the two daughter cells. Finally, we consider how extracellular signals govern the rates of cell growth and division and how these two processes are coordinated.


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Cell_Biology_Alberts. The most basic function of the cell cycle is to duplicate the vast amount of DNA in the chromosomes and then segregate the copies into two genetically identical daughter cells. These processes define the two major phases of the cell cycle. Chromosome duplication occurs during S phase (S for DNA synthesis), which requires 10–12 hours and occupies about half of the cell-cycle time in a typical mammalian cell. After S phase, chromosome segregation and cell division occur in M phase (M for mitosis), which requires much less time (less than an hour in a mammalian cell). M phase comprises two major events: nuclear division, or mitosis, during which the copied chromosomes are distributed into a pair of daughter nuclei; and cytoplasmic division, or cytokinesis, when the cell itself divides in two (Figure 17–2).


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Cell_Biology_Alberts. At the end of S phase, the DNA molecules in each pair of duplicated chromosomes are intertwined and held tightly together by specialized protein linkages. Early in mitosis at a stage called prophase, the two DNA molecules are gradually disentangled and condensed into pairs of rigid, compact rods called sister chromatids, which remain linked by sister-chromatid cohesion. When the nuclear envelope disassembles later in mitosis, the sister-chromatid pairs become attached to the mitotic spindle, a giant bipolar array of microtubules (discussed in Chapter 16). Sister chromatids are attached to opposite poles of the spindle and, eventually, align at the spindle equator in a stage called metaphase. The destruction of sister-chromatid cohesion at the start of anaphase separates the sister chromatids, which are pulled to opposite poles of the spindle. The spindle is then disassembled, and the segregated chromosomes are packaged into separate nuclei at telophase. Cytokinesis then cleaves the


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Cell_Biology_Alberts. which are pulled to opposite poles of the spindle. The spindle is then disassembled, and the segregated chromosomes are packaged into separate nuclei at telophase. Cytokinesis then cleaves the cell in two, so that each daughter cell inherits one of the two nuclei (Figure 17–3).


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Cell_Biology_Alberts. The Eukaryotic Cell Cycle Usually Consists of Four phases Most cells require much more time to grow and double their mass of proteins and organelles than they require to duplicate their chromosomes and divide. Partly to allow time for growth, most cell cycles have gap phases—a G1 phase between M phase and S phase and a G2 phase between S phase and mitosis. Thus, the eukaryotic cell cycle is traditionally divided into four sequential phases: G1, S, G2, and M. G1, S, and G2 together are called interphase (Figure 17–4, and see Figure 17–3). In a typical human cell proliferating in culture, interphase might occupy 23 hours of a 24-hour cycle, with 1 hour for M phase. Cell growth occurs throughout the cell cycle, except during mitosis. The two gap phases are more than simple time delays to allow cell growth. They also provide time for the cell to monitor the internal and external environment


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Cell_Biology_Alberts. The two gap phases are more than simple time delays to allow cell growth. They also provide time for the cell to monitor the internal and external environment Figure 17–2 The major events of the cell cycle. The major chromosomal events of the cell cycle occur in s phase, when the chromosomes are duplicated, and m phase, when the duplicated chromosomes are segregated into a pair of daughter nuclei (in mitosis), after which the cell itself divides into two (cytokinesis). Figure 17–1 The cell cycle. The division of a hypothetical eukaryotic cell with two chromosomes (one red, and one black) is shown to illustrate how two genetically identical daughter cells are produced in each cycle. Each of the daughter cells will often continue to divide by going through additional cell cycles.


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Cell_Biology_Alberts. Figure 17–3 The events of eukaryotic cell division as seen under a microscope. The easily visible processes of nuclear division (mitosis) and cell division (cytokinesis), collectively called m phase, typically occupy only a small fraction of the cell cycle. The other, much longer, part of the cycle is known as interphase, which includes s phase and the gap phases (discussed in text). The five stages of mitosis are shown: an abrupt change in the biochemical state of the cell occurs at the transition from metaphase to anaphase. a cell can pause in metaphase before this transition point, but once it passes this point, the cell carries on to the end of mitosis and through cytokinesis into interphase.


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Cell_Biology_Alberts. to ensure that conditions are suitable and preparations are complete before the cell commits itself to the major upheavals of S phase and mitosis. The G1 phase is especially important in this respect. Its length can vary greatly depending on external conditions and extracellular signals from other cells. If extracellular conditions are unfavorable, for example, cells delay progress through G1 and may even enter a specialized resting state known as G0 (G zero), in which they can remain for days, weeks, or even years before resuming proliferation. Indeed, many cells remain permanently in G0 until they or the organism dies. If extracellular conditions are favorable and signals to grow and divide are present, cells in early G1 or G0 progress through a commitment point near the end of G1 known as Start (in yeasts) or the restriction point (in mammalian cells). We will use the term Start for both yeast and animal cells. After passing this point, cells are committed to DNA replication, even


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Cell_Biology_Alberts. as Start (in yeasts) or the restriction point (in mammalian cells). We will use the term Start for both yeast and animal cells. After passing this point, cells are committed to DNA replication, even if the extracellular signals that stimulate cell growth and division are removed.


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Cell_Biology_Alberts. Some features of the cell cycle, including the time required to complete certain events, vary greatly from one cell type to another, even in the same organism. The basic organization of the cycle, however, is essentially the same in all eukaryotic Figure 17–4 The four phases of the cell cycle. In most cells, gap phases separate the major events of s phase and m phase. g1 is the gap between m phase and s phase, while g2 is the gap between s phase and m phase.


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Cell_Biology_Alberts. Figure 17–5 Mammalian cells proliferating in culture. The cells in this scanning electron micrograph are rat fibroblasts. Cells at the lower left have rounded up and are in mitosis. (Courtesy of guenter albrecht-Buehler.) cells, and all eukaryotes appear to use similar machinery and control mechanisms to drive and regulate cell-cycle events. The proteins of the cell-cycle control system, for example, first appeared over a billion years ago. Remarkably, they have been so well conserved over the course of evolution that many of them function perfectly when transferred from a human cell to a yeast cell. We can therefore study the cell cycle and its regulation in a variety of organisms and use the findings from all of them to assemble a unified picture of how eukaryotic cells divide.


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Cell_Biology_Alberts. Several model organisms are used in the analysis of the eukaryotic cell cycle. The budding yeast Saccharomyces cerevisiae and the fission yeast Schizosaccharomyces pombe are simple eukaryotes in which powerful molecular and genetic approaches can be used to identify and characterize the genes and proteins that govern the fundamental features of cell division. The early embryos of certain animals, particularly those of the frog Xenopus laevis, are excellent tools for biochemical dissection of cell-cycle control mechanisms, while the fruit fly Drosophila melanogaster is useful for the genetic analysis of mechanisms underlying the control and coordination of cell growth and division in multicellular organisms. Cultured human cells provide an excellent system for the molecular and microscopic exploration of the complex processes by which our own cells divide.


Cell_Biology_Alberts

Cell_Biology_Alberts. How can we tell what stage a cell has reached in the cell cycle? One way is simply to look at living cells with a microscope. A glance at a population of mammalian cells proliferating in culture reveals that a fraction of the cells have rounded up and are in mitosis (Figure 17–5). Others can be observed in the process of cytokinesis. Similarly, looking at budding yeast cells under a microscope is very useful, because the size of the bud provides an indication of cell-cycle stage (Figure 17–6). We can gain additional clues about cell-cycle position by staining cells with DNA-binding fluorescent dyes (which reveal the condensation of chromosomes in mitosis) or with antibodies that recognize specific cell components such as the microtubules (revealing the mitotic spindle). S-phase cells can be identified in the microscope by supplying them with visualizable molecules that are incorporated into newly synthesized DNA, such as the artificial thymidine analog bromodeoxyuridine (BrdU); cell


Cell_Biology_Alberts

Cell_Biology_Alberts. be identified in the microscope by supplying them with visualizable molecules that are incorporated into newly synthesized DNA, such as the artificial thymidine analog bromodeoxyuridine (BrdU); cell nuclei that have incorporated BrdU are then revealed by staining with anti-BrdU antibodies (Figure 17–7).


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Cell_Biology_Alberts. Typically, in a population of cultured mammalian cells that are all proliferating rapidly but asynchronously, about 30–40% will be in S phase at any instant and become labeled by a brief pulse of BrdU. From the proportion of cells in such a population that are labeled, we can estimate the duration of S phase as a fraction of the whole cell-cycle duration. Similarly, from the proportion of cells in mitosis (the mitotic index), we can estimate the duration of M phase. Another way to assess the stage that a cell has reached in the cell cycle is by measuring its DNA content, which doubles during S phase. This approach is greatly facilitated by the use of fluorescent DNA-binding dyes and a flow cytometer, which allows the rapid and automatic analysis of large numbers of cells (Figure 17–8). We can use flow cytometry to determine the lengths of G1, S, and G2 + M phases, by measuring DNA content in a synchronized cell population as it progresses through the cell cycle.


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Cell_Biology_Alberts. Figure 17–6 The morphology of budding yeast cells. In a normal population of proliferating yeast cells, buds vary in size according to the cell-cycle stage. Unbudded cells are in g1. progression through the start transition triggers formation of a tiny bud, which grows in size during the s and m phases until it is almost the size of the mother cell. (Courtesy of Jeff Ubersax.) 20 µm Figure 17–7 Labeling S-phase cells. an immunofluorescence micrograph of BrdU-labeled epithelial cells of the zebrafish gut. The fish was exposed to BrdU, after which the tissue was fixed and prepared for labeling with fluorescent anti-BrdU antibodies (green). all the cells are stained with a red fluorescent dye. (Courtesy of Cécile Crosnier.)


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Cell_Biology_Alberts. Cell division usually begins with duplication of the cell’s contents, followed by distribution of those contents into two daughter cells. Chromosome duplication occurs during S phase of the cell cycle, whereas most other cell components are duplicated continuously throughout the cycle. During M phase, the replicated chromosomes are segregated into individual nuclei (mitosis), and the cell then splits in two (cytokinesis). S phase and M phase are usually separated by gap phases called G1 and G2, when various intracellular and extracellular signals regulate cell-cycle progression. Cell-cycle organization and control have been highly conserved during evolution, and studies in a wide range of systems have led to a unified view of eukaryotic cell-cycle control.


Cell_Biology_Alberts

Cell_Biology_Alberts. For many years, cell biologists watched the puppet show of DNA synthesis, mitosis, and cytokinesis but had no idea of what lay behind the curtain controlling these events. It was not even clear whether there was a separate control system, or whether the processes of DNA synthesis, mitosis, and cytokinesis somehow controlled themselves. A major breakthrough came in the late 1980s with the identification of the key proteins of the control system, along with the realization that they are distinct from the proteins that perform the processes of DNA replication, chromosome segregation, and so on. In this section, we first consider the basic principles upon which the cell-cycle control system operates. We then discuss the protein components of the system and how they work together to time and coordinate the events of the cell cycle. The Cell-Cycle Control system Triggers the major Events of the Cell Cycle


Cell_Biology_Alberts

Cell_Biology_Alberts. The cell-cycle control system operates much like a timer that triggers the events of the cell cycle in a set sequence (Figure 17–9). In its simplest form—as seen in the stripped-down cell cycles of early animal embryos, for example—the control system is rigidly programmed to provide a fixed amount of time for the completion of each cell-cycle event. The control system in these early embryonic divisions is independent of the events it controls, so that its timing mechanisms continue to operate even if those events fail. In most cells, however, the control system does respond to information received back from the processes it controls. If some malfunction prevents the successful completion of DNA synthesis, for example, signals are sent to the control system to delay progression to M phase. Such delays provide time for the machinery to be repaired and also prevent the disaster that might result if the cycle progressed prematurely to the next stage—and segregated incompletely replicated


Cell_Biology_Alberts

Cell_Biology_Alberts. Such delays provide time for the machinery to be repaired and also prevent the disaster that might result if the cycle progressed prematurely to the next stage—and segregated incompletely replicated chromosomes, for example.


Cell_Biology_Alberts

Cell_Biology_Alberts. The cell-cycle control system is based on a connected series of biochemical switches, each of which initiates a specific cell-cycle event. This system of switches possesses many important features that increase the accuracy and reliability of cell-cycle progression. First, the switches are generally binary (on/off ) and launch events in a complete, irreversible fashion. It would clearly be disastrous, for example, if events like chromosome condensation or nuclear-envelope breakdown were only partially initiated or started but not completed. Second, the cell-cycle control system is remarkably robust and reliable, partly because backup mechanisms and other features allow the system to operate effectively under a variety of conditions and even if some components fail. Finally, the control system is highly adaptable and can be modified to suit specific cell types or to respond to specific intracellular or extracellular signals. number of cells 012 relative amount of DNA per cell


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Cell_Biology_Alberts. Figure 17–8 Analysis of DNA content with a flow cytometer. This graph shows typical results obtained for a proliferating cell population when the dna content of its individual cells is determined in a flow cytometer. (a flow cytometer, also called a fluorescence-activated cell sorter, or FaCs, can also be used to sort cells according to their fluorescence—see Figure 8–2). The cells analyzed here were stained with a dye that becomes fluorescent when it binds to dna, so that the amount of fluorescence is directly proportional to the amount of dna in each cell. The cells fall into three categories: those that have an unreplicated complement of dna and are therefore in g1, those that have a fully replicated complement of dna (twice the g1 dna content) and are in g2 or m phase, and those that have an intermediate amount of dna and are in s phase. The distribution of cells indicates that there are greater numbers of cells in g1 than in g2 + m phase, showing that g1 is longer than g2 + m in


Cell_Biology_Alberts

Cell_Biology_Alberts. have an intermediate amount of dna and are in s phase. The distribution of cells indicates that there are greater numbers of cells in g1 than in g2 + m phase, showing that g1 is longer than g2 + m in this population.


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Cell_Biology_Alberts. Are all chromosomes attached to the spindle? Is all DNA replicated? Is environment favorable? G2 M S G1 CONTROLLER ENTER MITOSIS TRIGGER ANAPHASE AND PROCEED TO CYTOKINESIS ENTER CELL CYCLE AND PROCEED TO S PHASE START TRANSITION Is environment favorable? G2/M TRANSITION


Cell_Biology_Alberts

Cell_Biology_Alberts. In most eukaryotic cells, the cell-cycle control system governs cell-cycle progression at three major regulatory transitions (see Figure 17–9). The first is Start (or the restriction point) in late G1, where the cell commits to cell-cycle entry and chromosome duplication. The second is the G2/M transition, where the control system triggers the early mitotic events that lead to chromosome alignment on the mitotic spindle in metaphase. The third is the metaphase-to-anaphase transition, where the control system stimulates sister-chromatid separation, leading to the completion of mitosis and cytokinesis. The control system blocks progression through each of these transitions if it detects problems inside or outside the cell. If the control system senses problems in the completion of DNA replication, for example, it will hold the cell at the G2/M transition until those problems are solved. Similarly, if extracellular conditions are not appropriate for cell proliferation, the control system


Cell_Biology_Alberts

Cell_Biology_Alberts. for example, it will hold the cell at the G2/M transition until those problems are solved. Similarly, if extracellular conditions are not appropriate for cell proliferation, the control system blocks progression through Start, thereby preventing cell division until conditions become favorable.


Cell_Biology_Alberts

Cell_Biology_Alberts. The Cell-Cycle Control system depends on Cyclically activated Cyclin-dependent protein kinases (Cdks) Central components of the cell-cycle control system are members of a family of protein kinases known as cyclin-dependent kinases (Cdks). The activities of these kinases rise and fall as the cell progresses through the cycle, leading to cyclical changes in the phosphorylation of intracellular proteins that initiate or regulate the major events of the cell cycle. An increase in Cdk activity at the G2/M transition, for example, increases the phosphorylation of proteins that control chromosome condensation, nuclear-envelope breakdown, spindle assembly, and other events that occur in early mitosis.


Cell_Biology_Alberts

Cell_Biology_Alberts. Cyclical changes in Cdk activity are controlled by a complex array of enzymes and other proteins. The most important of these Cdk regulators are proteins known as cyclins. Cdks, as their name implies, are dependent on cyclins for their activity: unless they are bound tightly to a cyclin, they have no protein kinase activity (Figure 17–10). Cyclins were originally named because they undergo a cycle of synthesis and degradation in each cell cycle. The levels of the Cdk proteins, by contrast, are constant. Cyclical changes in cyclin protein levels result in the cyclic assembly and activation of cyclin–Cdk complexes at specific stages of the cell cycle. Figure 17–9 The control of the cell cycle.


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Cell_Biology_Alberts. Figure 17–9 The control of the cell cycle. a cell-cycle control system triggers the essential processes of the cycle—such as dna replication, mitosis, and cytokinesis. The control system is represented here as a central arm—the controller—that rotates clockwise, triggering essential processes when it reaches specific transitions on the outer dial (yellow boxes). Information about the completion of cell-cycle events, as well as signals from the environment, can cause the control system to arrest the cycle at these transitions. Figure 17–10 Two key components of the cell-cycle control system. When cyclin forms a complex with Cdk, the protein kinase is activated to trigger specific cell-cycle events. Without cyclin, Cdk is inactive. There are four classes of cyclins, each defined by the stage of the cell cycle at which they bind Cdks and function. All eukaryotic cells require three of these classes (Figure 17–11): 1.


Cell_Biology_Alberts

Cell_Biology_Alberts. There are four classes of cyclins, each defined by the stage of the cell cycle at which they bind Cdks and function. All eukaryotic cells require three of these classes (Figure 17–11): 1. G1/S-cyclins activate Cdks in late G1 and thereby help trigger progression through Start, resulting in a commitment to cell-cycle entry. Their levels fall in S phase. 2. S-cyclins bind Cdks soon after progression through Start and help stimulate chromosome duplication. S-cyclin levels remain elevated until mitosis, and these cyclins also contribute to the control of some early mitotic events. 3. M-cyclins activate Cdks that stimulate entry into mitosis at the G2/M transition. M-cyclin levels fall in mid-mitosis. In most cells, a fourth class of cyclins, the G1-cyclins, helps govern the activities of the G1/S-cyclins, which control progression through Start in late G1.


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Cell_Biology_Alberts. In yeast cells, a single Cdk protein binds all classes of cyclins and triggers different cell-cycle events by changing cyclin partners at different stages of the cycle. In vertebrate cells, by contrast, there are four Cdks. Two interact with G1-cyclins, one with G1/Sand S-cyclins, and one with Sand M-cyclins. In this chapter, we simply refer to the different cyclin–Cdk complexes as G1-Cdk, G1/S-Cdk, S-Cdk, and M-Cdk. Table 17–1 lists the names of the individual Cdks and cyclins.


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Cell_Biology_Alberts. How do different cyclin–Cdk complexes trigger different cell-cycle events? The answer, at least in part, seems to be that the cyclin protein does not simply activate its Cdk partner but also directs it to specific target proteins. As a result, each cyclin–Cdk complex phosphorylates a different set of substrate proteins. The same cyclin–Cdk complex can also induce different effects at different times in the cycle, probably because the accessibility of some Cdk substrates changes during the cell cycle. Certain proteins that function in mitosis, for example, may become available for phosphorylation only in G2.


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Cell_Biology_Alberts. Figure 17–11 Cyclin–Cdk complexes of the cell-cycle control system. The concentrations of the three major cyclin types oscillate during the cell cycle, while the concentrations of Cdks (not shown) do not change and exceed cyclin amounts. In late g1, rising g1/s-cyclin levels lead to the formation of g1/s-Cdk complexes that trigger progression through the start transition. s-Cdk complexes form at the start of s phase and trigger dna replication, as well as some early mitotic events. m-Cdk complexes form during g2 but are held in an inactive state; they are activated at the end of g2 and trigger entry into mitosis at the g2/m transition. a separate regulatory protein complex, the apC/C, initiates the metaphase-toanaphase transition, as we discuss later.


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Cell_Biology_Alberts. Studies of the three-dimensional structures of Cdk and cyclin proteins have revealed that, in the absence of cyclin, the active site in the Cdk protein is partly obscured by a protein loop, like a stone blocking the entrance to a cave (Figure 17–12A). Cyclin binding causes the loop to move away from the active site, resulting in partial activation of the Cdk enzyme (Figure 17–12B). Full activation of the cyclin–Cdk complex then occurs when a separate kinase, the Cdk-activating kinase (CAK), phosphorylates an amino acid near the entrance of the Cdk active site. This causes a small conformational change that further increases the activity of the Cdk, allowing the kinase to phosphorylate its target proteins effectively and thereby induce specific cell-cycle events (Figure 17–12C). The rise and fall of cyclin levels is the primary determinant of Cdk activity during the cell cycle. Several additional mechanisms, however, help control Cdk activity at specific stages of the cycle.


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Cell_Biology_Alberts. Phosphorylation at a pair of amino acids in the roof of the kinase active site inhibits the activity of a cyclin–Cdk complex. Phosphorylation of these sites by a protein kinase known as Wee1 inhibits Cdk activity, while dephosphorylation of these sites by a phosphatase known as Cdc25 increases Cdk activity (Figure 17–13). We will see later that this regulatory mechanism is particularly important in the control of M-Cdk activity at the onset of mitosis. Binding of Cdk inhibitor proteins (CKIs) inactivates cyclin–Cdk complexes. The three-dimensional structure of a cyclin–Cdk–CKI complex reveals that CKI binding stimulates a large rearrangement in the structure of the Cdk active site, rendering it inactive (Figure 17–14). Cells use CKIs primarily to help govern the activities of G1/Sand S-Cdks early in the cell cycle. Regulated proteolysis Triggers the metaphase-to-anaphase Transition


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Cell_Biology_Alberts. Regulated proteolysis Triggers the metaphase-to-anaphase Transition Whereas activation of specific cyclin–Cdk complexes drives progression through the Start and G2/M transitions (see Figure 17–11), progression through the meta-phase-to-anaphase transition is triggered not by protein phosphorylation but by protein destruction, leading to the final stages of cell division. The key regulator of the metaphase-to-anaphase transition is the anaphasepromoting complex, or cyclosome (APC/C), a member of the ubiquitin ligase family of enzymes. As discussed in Chapter 3, these enzymes are used in numerous cell processes to stimulate the proteolytic destruction of specific regulatory proteins. They polyubiquitylate specific target proteins, resulting in their destruction in proteasomes. Other ubiquitin ligases mark proteins for purposes other than destruction (discussed in Chapter 3).


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Cell_Biology_Alberts. Figure 17–12 The structural basis of Cdk activation. These drawings are based on three-dimensional structures of human Cdk2 and cyclin a, as determined by x-ray crystallography. The location of the bound aTp is indicated. The enzyme is shown in three states. (a) In the inactive state, without cyclin bound, the active site is blocked by a region of the protein called the T-loop (red). (B) The binding of cyclin causes the T-loop to move out of the active site, resulting in partial activation of the Cdk2. (C) phosphorylation of Cdk2 (by Cak) at a threonine residue in the T-loop further activates the enzyme by changing the shape of the T-loop, improving the ability of the enzyme to bind its protein substrates (Movie 17.1).


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Cell_Biology_Alberts. Figure 17–13 The regulation of Cdk activity by phosphorylation. The active cyclin–Cdk complex is turned off when the kinase Wee1 phosphorylates two closely spaced sites above the active site. Removal of these phosphates by the phosphatase Cdc25 activates the cyclin– Cdk complex. For simplicity, only one inhibitory phosphate is shown. Cak adds the activating phosphate, as shown in Figure 17–12. Figure 17–14 The inhibition of a cyclin–Cdk complex by a CKI. This drawing is based on the three-dimensional structure of the human cyclin a–Cdk2 complex bound to the CkI p27, as determined by x-ray crystallography. The p27 binds to both the cyclin and Cdk in the complex, distorting the active site of the Cdk. It also inserts into the aTp-binding site, further inhibiting the enzyme activity.


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Cell_Biology_Alberts. The APC/C catalyzes the ubiquitylation and destruction of two major types of proteins. The first is securin, which protects the protein linkages that hold sister-chromatid pairs together in early mitosis. Destruction of securin in meta-phase activates a protease that separates the sisters and unleashes anaphase, as described later. The Sand M-cyclins are the second major targets of the APC/C. Destroying these cyclins inactivates most Cdks in the cell (see Figure 17–11). As a result, the many proteins phosphorylated by Cdks from S phase to early mitosis are dephosphorylated by various phosphatases in the anaphase cell. This dephosphorylation of Cdk targets is required for the completion of M phase, including the final steps in mitosis and then cytokinesis. Following its activation in mid-mitosis, the APC/C remains active in G1 to provide a stable period of Cdk inactivity. When G1/S-Cdk is activated in late G1, the APC/C is turned off, thereby allowing cyclin accumulation in the next


Cell_Biology_Alberts

Cell_Biology_Alberts. the APC/C remains active in G1 to provide a stable period of Cdk inactivity. When G1/S-Cdk is activated in late G1, the APC/C is turned off, thereby allowing cyclin accumulation in the next cell cycle.


Cell_Biology_Alberts

Cell_Biology_Alberts. The cell-cycle control system also uses another ubiquitin ligase called SCF (see Figure 3–71). It has many functions in the cell, but its major role in the cell cycle is to ubiquitylate certain CKI proteins in late G1, thereby helping to control the activation of S-Cdks and DNA replication. SCF is also responsible for the destruction of G1/S-cyclins in early S phase.


Cell_Biology_Alberts

Cell_Biology_Alberts. The APC/C and SCF are both large, multisubunit complexes with some related components (see Figure 3–71), but they are regulated differently. APC/C activity changes during the cell cycle, primarily as a result of changes in its association with an activating subunit—either Cdc20 in mid-mitosis or Cdh1 from late mitosis through early G1. These subunits help the APC/C recognize its target proteins (Figure 17–15A). SCF activity depends on substrate-binding subunits called F-box proteins. Unlike APC/C activity, however, SCF activity is constant during the cell cycle. Ubiquitylation by SCF is controlled instead by changes in the phosphorylation state of its target proteins, as F-box subunits recognize only specifically phosphorylated proteins (Figure 17–15B).


Cell_Biology_Alberts

Cell_Biology_Alberts. In the simple cell cycles of early animal embryos, gene transcription does not occur. Cell-cycle control depends exclusively on post-transcriptional mechanisms that involve the regulation of Cdks and ubiquitin ligases and their target proteins. In the more complex cell cycles of most cell types, however, transcriptional control provides an important additional level of regulation. Changes in cyclin gene transcription, for example, help control cyclin levels in most cells. A variety of methods discussed in Chapter 8 have been used to analyze changes in the expression of all genes in the genome as the cell progresses through the cell cycle. The results of these studies are surprising. In budding yeast, for example, about 10% of the genes encode mRNAs whose levels oscillate during the cell cycle. Some of these genes encode proteins with known cell-cycle functions, but the functions of many others are unknown. The Cell-Cycle Control system Functions as a network of Biochemical switches


Cell_Biology_Alberts

Cell_Biology_Alberts. The Cell-Cycle Control system Functions as a network of Biochemical switches Table 17–2 summarizes some of the major components of the cell-cycle control system. These proteins are functionally linked to form a robust network, which operates essentially autonomously to activate a series of biochemical switches, each of which triggers a specific cell-cycle event. When conditions for cell proliferation are right, various external and internal signals stimulate the activation of G1-Cdk, which in turn stimulates the expression of genes encoding G1/Sand S-cyclins (Figure 17–16). The resulting activation of G1/S-Cdk then drives progression through the Start transition. By mechanisms we discuss later, G1/S-Cdks unleash a wave of S-Cdk activity, which initiates (A) control of proteolysis by APC/C (B) control of proteolysis by SCF


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Cell_Biology_Alberts. Figure 17–15 The control of proteolysis by APC/C and SCF during the cell cycle. (a) The apC/C is activated in mitosis by association with Cdc20, which recognizes specific amino acid sequences on m-cyclin and other target proteins. With the help of two additional proteins called E1 and E2, the apC/C assembles polyubiquitin chains on the target protein. The polyubiquitylated target is then recognized and degraded in a proteasome. (B) The activity of the ubiquitin ligase sCF depends on substrate-binding subunits called F-box proteins, of which there are many different types. The phosphorylation of a target protein, such as the CkI shown, allows the target to be recognized by a specific F-box subunit.


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Cell_Biology_Alberts. chromosome duplication in S phase and also contributes to some early events of mitosis. M-Cdk activation then triggers progression through the G2/M transition and the events of early mitosis, leading to the alignment of sister-chromatid pairs at the equator of the mitotic spindle. Finally, the APC/C, together with its activator Cdc20, triggers the destruction of securin and cyclins, thereby unleashing sister-chromatid separation and segregation and the completion of mitosis. When mitosis is complete, multiple mechanisms collaborate to suppress Cdk activity, resulting in a stable G1 period. We are now ready to discuss these cell-cycle stages in more detail, starting with S phase.


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Cell_Biology_Alberts. Figure 17–16 An overview of the cell-cycle control system. The core of the cell-cycle control system consists of a series of cyclin–Cdk complexes (yellow). The activity of each complex is also influenced by various inhibitory mechanisms, which provide information about the extracellular environment, cell damage, and incomplete cell-cycle events (top). These inhibitory mechanisms are not present in all cell types; many are missing in early embryonic cell cycles, for example. The cell-cycle control system triggers the events of the cell cycle and ensures that they are properly timed and coordinated with each other. The control system responds to various intracellular and extracellular signals and arrests the cycle when the cell either fails to complete an essential cell-cycle process or encounters unfavorable environmental or intracellular conditions.


Cell_Biology_Alberts

Cell_Biology_Alberts. Central components of the control system are the cyclin-dependent protein kinases (Cdks), which depend on cyclin subunits for their activity. Oscillations in the activities of different cyclin–Cdk complexes control various cell-cycle events. Thus, activation of S-phase cyclin–Cdk complexes (S-Cdk) initiates S phase, whereas activation of M-phase cyclin–Cdk complexes (M-Cdk) triggers mitosis. The mechanisms that control the activities of cyclin–Cdk complexes include phosphorylation of the Cdk subunit, binding of Cdk inhibitor proteins (CKIs), proteolysis of cyclins, and changes in the transcription of genes encoding Cdk regulators. The cell-cycle control system also depends crucially on two additional enzyme complexes, the APC/C and SCF ubiquitin ligases, which catalyze the ubiquitylation and consequent destruction of specific regulatory proteins that control critical events in the cycle.


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Cell_Biology_Alberts. The linear chromosomes of eukaryotic cells are vast and dynamic assemblies of DNA and protein, and their duplication is a complex process that takes up a major fraction of the cell cycle. Not only must the long DNA molecule of each chromosome be duplicated accurately—a remarkable feat in itself—but the protein packaging surrounding each region of that DNA must also be reproduced, ensuring that the daughter cells inherit all features of chromosome structure.


Cell_Biology_Alberts

Cell_Biology_Alberts. The central event of chromosome duplication—DNA replication—poses two problems for the cell. First, replication must occur with extreme accuracy to minimize the risk of mutations in the next cell generation. Second, every nucleotide in the genome must be copied once, and only once, to prevent the damaging effects of gene amplification. In Chapter 5, we discuss the sophisticated protein machinery that performs DNA replication with astonishing speed and accuracy. In this section, we consider the elegant mechanisms by which the cell-cycle control system initiates the replication process and, at the same time, prevents it from happening more than once per cycle.


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Cell_Biology_Alberts. DNA replication begins at origins of replication, which are scattered at numerous locations in every chromosome. During S phase, DNA replication is initiated at these origins when a DNA helicase unwinds the double helix and DNA replication enzymes are loaded onto the two single-stranded templates. This leads to the elongation phase of replication, when the replication machinery moves outward from the origin at two replication forks (discussed in Chapter 5).


Cell_Biology_Alberts

Cell_Biology_Alberts. To ensure that chromosome duplication occurs only once per cell cycle, the initiation phase of DNA replication is divided into two distinct steps that occur at different times in the cell cycle (Figure 17–17). The first step occurs in late mitosis and early G1, when a pair of inactive DNA helicases is loaded onto the replication origin, forming a large complex called the prereplicative complex or preRC. This step is sometimes called licensing of replication origins because initiation of DNA synthesis is permitted only at origins containing a preRC. The second step occurs in S phase, when the DNA helicases are activated, resulting in DNA unwinding and the initiation of DNA synthesis. Once a replication origin has been fired in this way, the two helicases move out from the origin with the replication forks, and that origin cannot be reused until a new preRC is assembled there at the end of mitosis. As a result, origins can be activated only once per cell cycle.


Cell_Biology_Alberts

Cell_Biology_Alberts. Figure 17–18 illustrates some of the molecular details underlying the control of the two steps in the initiation of DNA replication. A key player is a large multiprotein complex called the origin recognition complex (ORC), which binds to Figure 17–17 Control of chromosome duplication. preparations for dna replication begin in late mitosis and g1, when the dna helicases are loaded by multiple proteins at the replication origin, forming the prereplicative complex (preRC). s-Cdk activation leads to activation of the dna helicases, which unwind the dna at origins to initiate dna replication. Two replication forks move out from each origin until the entire chromosome is duplicated. duplicated chromosomes are then segregated in m phase. s-Cdk activation in s phase also prevents assembly of new preRCs at any origin until the following g1—thereby ensuring that each origin is activated only once in each cell cycle.


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Cell_Biology_Alberts. replication origins throughout the cell cycle. In late mitosis and early G1, the proteins Cdc6 and Cdt1 collaborate with the ORC to load the inactive DNA helicases around the DNA next to the origin. The resulting large complex is the preRC, and the origin is now licensed for replication. At the onset of S phase, S-Cdk triggers origin activation by phosphorylating specific initiator proteins, which then nucleate the assembly of a large protein complex that activates the DNA helicase and recruits the DNA synthesis machinery. Another protein kinase called DDK is also activated in S phase and helps drive origin activation by phosphorylating specific subunits of the DNA helicase.


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Cell_Biology_Alberts. At the same time as S-Cdk initiates DNA replication, several mechanisms prevent assembly of new preRCs. S-Cdk phosphorylates and thereby inhibits the ORC and Cdc6 proteins. Inactivation of the APC/C in late G1 also helps turn off preRC assembly. In late mitosis and early G1, the APC/C triggers the destruction of a Cdt1 inhibitor called geminin, thereby allowing Cdt1 to be active. When the APC/C is turned off in late G1, geminin accumulates and inhibits the Cdt1 that is not associated with DNA. Also, the association of Cdt1 with a protein at active replication forks stimulates Cdt1 destruction. In these various ways, preRC formation is prevented from S phase to mitosis, thereby ensuring that each origin is fired only once per cell cycle. How, then, is the cell-cycle control system reset to allow replication in the next cell cycle? At the end of mitosis, APC/C activation leads to the inactivation of Cdks and the destruction of geminin. ORC and Cdc6 are dephosphorylated and Cdt1 is


Cell_Biology_Alberts

Cell_Biology_Alberts. to allow replication in the next cell cycle? At the end of mitosis, APC/C activation leads to the inactivation of Cdks and the destruction of geminin. ORC and Cdc6 are dephosphorylated and Cdt1 is activated, allowing preRC assembly to prepare the cell for the next S phase.


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Cell_Biology_Alberts. Chromosome duplication Requires duplication of Chromatin structure The DNA of the chromosomes is extensively packaged in a variety of protein components, including histones and various regulatory proteins involved in the control of gene expression (discussed in Chapter 4). Thus, duplication of a chromosome is not simply a matter of replicating the DNA at its core but also requires the duplication of these chromatin proteins and their proper assembly on the DNA.


Cell_Biology_Alberts

Cell_Biology_Alberts. The production of chromatin proteins increases during S phase to provide the raw materials needed to package the newly synthesized DNA. Most importantly, S-Cdks stimulate a large increase in the synthesis of the four histone subunits that form the histone octamers at the core of each nucleosome. These subunits are assembled into nucleosomes on the DNA by nucleosome assembly factors, which typically associate with the replication fork and distribute nucleosomes on both strands of the DNA as they emerge from the DNA synthesis machinery.


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Cell_Biology_Alberts. Chromatin packaging helps to control gene expression. In some parts of the chromosome, the chromatin is highly condensed and is called heterochromatin, whereas in other regions it has a more open structure and is called euchromatin (discussed in Chapter 4). These differences in chromatin structure depend on a variety of mechanisms, including modification of histone tails and the presence of non-histone proteins. Because these differences are important in gene regulation, Cdc6 ORC (origin recognition complex) P P P P P P P P originCdt1 Mcm helicase + prereplicative complex (preRC) initiator proteins S-Cdk DDK INACTIVATION OF ORC, Cdc6, Cdt1 DNA HELICASE ACTIVATION COMPLETION OF DNA REPLICATION G1 S G2 DNA


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Cell_Biology_Alberts. Figure 17–18 Control of the initiation of DNA replication. The replication origin is bound by the ORC throughout the cell cycle. In early g1, Cdc6 associates with the ORC, and these proteins bind the dna helicase, which contains six closely related subunits called mcm proteins. The helicase also associates with a protein called Cdt1. Using energy provided by aTp hydrolysis, the ORC and Cdc6 proteins load two copies of the dna helicase, in an inactive form, around the dna next to the origin, thereby forming the prereplicative complex (preRC). at the onset of s phase, s-Cdk stimulates the assembly of several initiator proteins on each dna helicase, while another protein kinase, ddk, phosphorylates subunits of the dna helicase. as a result, the dna helicases are activated and unwind the dna. dna polymerase and other replication proteins are recruited to the origin, and dna replication begins. The ORC is displaced by the replication machinery and then rebinds. s-Cdk and other mechanisms


Cell_Biology_Alberts

Cell_Biology_Alberts. dna polymerase and other replication proteins are recruited to the origin, and dna replication begins. The ORC is displaced by the replication machinery and then rebinds. s-Cdk and other mechanisms also inactivate the preRC components ORC, Cdc6, and Cdt1, thereby preventing formation of new preRCs at the origins until the end of mitosis (see text).


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Cell_Biology_Alberts. it is crucial that chromatin structure, like the DNA within, is reproduced accurately during S phase. How chromatin structure is reproduced is not well understood, however. During DNA synthesis, histone-modifying enzymes and various non-histone proteins are probably deposited onto the two new DNA strands as they emerge from the replication fork, and these proteins are thought to reproduce the local chromatin structure of the parent chromosome (see Figure 4–45).


Cell_Biology_Alberts

Cell_Biology_Alberts. At the end of S phase, each replicated chromosome consists of a pair of identical sister chromatids glued together along their length. This sister-chromatid cohesion sets the stage for a successful mitosis because it greatly facilitates the attachment of the two sister chromatids to opposite poles of the mitotic spindle. Imagine how difficult it would be to achieve this bipolar attachment if sister chromatids were allowed to drift apart after S phase. Indeed, defects in sister-chromatid cohesion—in yeast mutants, for example—lead inevitably to major errors in chromosome segregation.


Cell_Biology_Alberts

Cell_Biology_Alberts. Sister-chromatid cohesion depends on a large protein complex called cohesin, which is deposited at many locations along the length of each sister chromatid as the DNA is replicated in S phase. Two of the subunits of cohesin are members of a large family of proteins called SMC proteins (for Structural Maintenance of Chromosomes). Cohesin forms giant ringlike structures, and it has been proposed that these surround the two sister chromatids (Figure 17–19).


Cell_Biology_Alberts

Cell_Biology_Alberts. Sister-chromatid cohesion also results, at least in part, from DNA catenation, the intertwining of sister DNA molecules that occurs when two replication forks meet during DNA synthesis. The enzyme topoisomerase II gradually disentangles the catenated sister DNAs between S phase and early mitosis by cutting one DNA molecule, passing the other through the break, and then resealing the cut DNA (see Figure 5–22). Once the catenation has been removed, sister-chromatid cohesion depends primarily on cohesin complexes. The sudden and synchronous loss of sister cohesion at the metaphase-to-anaphase transition therefore depends primarily on disruption of these complexes, as we describe later.


Cell_Biology_Alberts

Cell_Biology_Alberts. Duplication of the chromosomes in S phase involves the accurate replication of the entire DNA molecule in each chromosome, as well as the duplication of the chromatin proteins that associate with the DNA and govern various aspects of chromosome function. Chromosome duplication is triggered by the activation of S-Cdk, which activates proteins that unwind the DNA and initiate its replication at replication origins. Once a replication origin is activated, S-Cdk also inhibits proteins that are required to allow that origin to initiate DNA replication again. Thus, each origin is fired once and only once in each S phase and cannot be reused until the next cell cycle.


Cell_Biology_Alberts

Cell_Biology_Alberts. Figure 17–19 Cohesin. Cohesin is a protein complex with four subunits. (a) Two subunits, smc1 and smc3, are coiled-coil proteins with an aTpase domain at one end; (B) two additional subunits, scc1 and scc3, connect the aTpase head domains, forming a ring structure that may encircle the sister chromatids as shown in (C). The aTpase domains are required for cohesin loading on the dna.


Cell_Biology_Alberts

Cell_Biology_Alberts. Following the completion of S phase and transition through G2, the cell undergoes the dramatic upheaval of M phase. This begins with mitosis, during which the sister chromatids are separated and distributed (segregated) to a pair of identical daughter nuclei, each with its own copy of the genome. Mitosis is traditionally divided into five stages—prophase, prometaphase, metaphase, anaphase, and telophase—defined primarily on the basis of chromosome behavior as seen in a microscope. As mitosis is completed, the second major event of M phase—cytokinesis—divides the cell into two halves, each with an identical nucleus. Panel 17–1 summarizes the major events of M phase (Movie 17.2, Movie 17.3, Movie 17.4, and Movie 17.5).


Cell_Biology_Alberts

Cell_Biology_Alberts. From a regulatory point of view, mitosis can be divided into two major parts, each governed by distinct components of the cell-cycle control system. First, an abrupt increase in M-Cdk activity at the G2/M transition triggers the events of early mitosis (prophase, prometaphase, and metaphase). During this period, M-Cdk and several other mitotic protein kinases phosphorylate a variety of proteins, leading to the assembly of the mitotic spindle and its attachment to the sister-chromatid pairs. The second major part of mitosis begins at the metaphaseto-anaphase transition, when the APC/C triggers the destruction of securin, liberating a protease that cleaves cohesin and thereby initiates separation of the sister chromatids. The APC/C also promotes the destruction of cyclins, which leads to Cdk inactivation and the dephosphorylation of Cdk targets, which is required for all events of late M phase, including the completion of anaphase, the disassembly of the mitotic spindle, and the


Cell_Biology_Alberts

Cell_Biology_Alberts. to Cdk inactivation and the dephosphorylation of Cdk targets, which is required for all events of late M phase, including the completion of anaphase, the disassembly of the mitotic spindle, and the division of the cell by cytokinesis.


Cell_Biology_Alberts

Cell_Biology_Alberts. In this section, we describe the key mechanical events of mitosis and how M-Cdk and the APC/C orchestrate them. One of the most remarkable features of cell-cycle control is that a single protein kinase, M-Cdk, brings about all of the diverse and complex cell rearrangements that occur in the early stages of mitosis. At a minimum, M-Cdk must induce the assembly of the mitotic spindle and ensure that each sister chromatid in a pair is attached to the opposite pole of the spindle. It also triggers chromosome condensation, the large-scale reorganization of the intertwined sister chromatids into compact, rodlike structures. In animal cells, M-Cdk also promotes the breakdown of the nuclear envelope and rearrangements of the actin cytoskeleton and the Golgi apparatus. Each of these processes is thought to be initiated when M-Cdk phosphorylates specific proteins involved in the process, although most of these proteins have not yet been identified.


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Cell_Biology_Alberts. M-Cdk does not act alone to phosphorylate key proteins involved in early mitosis. Two additional families of protein kinases, the Polo-like kinases and the Aurora kinases, also make important contributions to the control of early mitotic events. The Polo-like kinase Plk, for example, is required for the normal assembly of a bipolar mitotic spindle, in part because it phosphorylates proteins involved in separation of the spindle poles early in mitosis. The Aurora kinase Aurora-A also helps control proteins that govern the assembly and stability of the spindle, whereas Aurora-B controls attachment of sister chromatids to the spindle, as we discuss later. dephosphorylation activates m-Cdk at the Onset of mitosis