Lodish • Berk • Kaiser • Krieger • scott • Bretscher • Ploegh • Matsudaira

MOLECULAR CELL BIOLOGY SEVENTH EDITION

CHAPTER 18 Cell Organization and Movement II:

Microtubules and Intermediate Filaments

Copyr ight © 2013 by W. H. Freeman and Company

Figure 18.1 Overview of the physical properties and functions of the three cytoskeletal systems in animal cells.

Figure 18.3 Structure of tubulin dimers and their organization into microtubules.

The GTP bound to the alpha tubulin monomer is nonexchangeable, whereas the GDP bound to the beta tubulin monomer is exchangeable with GTP

Structure of tubulin dimers and their organization into microtubules 1. In the early days of electron microscopy, cell biologists noted long tubules in the cytoplasm that they called microtubules. Morphologically similar microtubules were seen making up the fibers of the mitotic spindle, as components of axons, and as the structural elements in cilia and flagella. Tubulin isolated in a pure and soluble form consists of two closely related subunits called α- and β-tubulin, each with a molecular weight of about 55,000 daltons. 2. The α- and β-subunits of the tubulin dimer can each bind one molecule of GTP (Figure 18-3a). The GTP in the α-tubulin subunit is never hydrolyzed and is trapped by the interface between the α- and β-subunits. By contrast, the GTP-binding site on the β-subunit is at the surface of the dimer. GTP bound by the β-subunit can be hydrolyzed, and the resulting GDP can be exchanged for free GTP. Under appropriate conditions, soluble tubulin dimers can assemble into microtubules (Figure 18-3b). As we saw in Chapter 17 for the polymerization of actin, ATP-G actin is preferentially added to one end of the filament, designated the (+) end because it is the end favored for assembly. Once incorporated into the filament, the bound ATP is hydrolyzed to ADP and Pi. In a similar manner, tubulin dimers in which the β-subunit has bound GTP add preferentially to one end of the microtubule, also designated the (+) end. As we will see, the GTP is hydrolyzed once tubulin is incorporated into the microtubule, but in contrast to the situation with ATP hydrolysis in an actin filament, this GTP hydrolysis has dramatic effects on the behavior of the microtubule (+) end. Microtubules are composed of 13 laterally associated protofilaments that form a tubule whose external diameter is about 25nm (see Figure 18-3b). Each of the 13 protofilaments is a string of αβ-tubulin dimers, longitudinally arranged so that the subunits alternate down a protofilament, with each subunit type repeating every 8nm. Because the αβ-tubulin dimers in a protofilament are all oriented in the same way, each protofilament has an α-subunit at one end and a β-subunit at the other-so the protofilaments have an intrinsic polarity.

Figure 18.4 Singlet, doublet, and triplet microtubules.

Singlet, doublet, and triplet microtubules Most microtubules in a cell consist of a simple tube, a singlet microtubule, built from 13 protofilaments. In rare cases, singlet microtubules contain more or fewer protofilaments; for example, certain microtubules in the neurons of nematode worms contain 11 or 15 protofilaments. In addition to the simple singlet structure, doublet or triplet microtubules are found in specialized structures such as cilia and flagella (doublet microtubules) and centrioles and basal bodies (triplet microtubules), structures we will explore later in the chapter. Each doublet or triplet contains one complete 13-protofilament microtubule (called the A tubule) and one or two additional tubules (B and C) consisting of 10 protofilaments each (Figure 18-4).

Figure 18.5 Microtubules are assembled from microtubule organizing centers (MTOCs).

Microtubules are assembled from microtubule organizing centers (MTOCs) The nucleation phase of microtubule assembly is such an unfavorable reaction that spontaneous nucleation does not play a significant role in microtubule assembly in vivo. Rather, all microtubules are nucleated from structures known as microtubule-organizing centers, or MTOCs. In most cases the (-) end of the microtubule stays anchored in the MTOC while the (+) end extends away from it. The centrosome is the main MTOC in animal cells. In nonmitotic cells, also known as interphase cells, the centrosome is generally located near the nucleus, producing an array of microtubules with their (+) ends radiating toward the cell periphery (Figure 18-5c). This radial display provides tracks for microtubule-based motor proteins to organize and transport membrane-bound compartments, such as those comprising the secretory and endocytic pathways. During mitosis, cells completely reorganize their microtubules to form a bipolar spindle, assembled from two centrosomes, also known as spindle poles, to accurately segregate copies of the duplicated chromosomes (Figrue 18-5d). In another example, neurons have long processes called axons, in which organelles are transported in both directions along microtubules (Figure 18-5e). The microtubules in axons, which can be as long as 1 meter in length, are not continuous and have been released from the centrosome but nevertheless are all of the same polarity. In the same cells, the microtubules in the dendrites have mixed polarity, although the funtional significance of this is not clear. In cilia and flagella (Figure 18-5f), microtubules are assembled from an MTOC called a basal body. As we mention later, plants do not have centrosomes and basal bodies but use other mechanisms to nucleate the assembly of microtubules.

Figure 18.6 Structure of centrosomes.

Figure 18.7 The γ-tubulin ring complex (γ-TuRC) that nucleates microtubule assembly.

Figure 16-30b Molecular Biology of the Cell (© Garland Science 2008)

Centrosomes 1. Electron microscopy shows that centrosomes in animal cells consist of a pair of orthogonally arranged cylindrical centrioles surrounded by apparently amorphous material called pericentriolar material (Figure 18-6a, arrowheads). Centrioles, which are about 0.5um long and 0.2um in diameter, are highly organized and stable structures that consist of nine sets of triplet microtubules and are closely related in structure to the basal bodies found at the base of cilia and flagella. It is not the centrioles themselves that nucleate the cytoplasmic microtubule array, but rather factors in the pericentriolar material. A critical component is the γ-tubulin ring complex (γ-TuRC) (Figure 18-6b and 18-7). γ-TuRC is located in the pericentriolar material and consists of many copies of γ-tubulin associated with several other proteins. It is believed that γ-TuRC acts like a split-washer template to bind αβ-tubulin dimers for the formation of a new microtubule, with the (-) end associated with γ-TuRC and the (+) end free for assembly. In addition to nucleating the assembly of microtubules, centrosomes anchor and regulate the dynamics of the (-) ends of the microtubules, which are located there. 2. Embedded in the centrosome is a pair of cylindrical structures arranged at right angles to each other in an L-shaped configuration (Figure 16-30b Molecular Biology of the Cell). These are the centrioles, which become the basal bodies of cilia and flagella in motile cells. The centrioles organize the centrosome matrix (also called the pericentriolar material), ensuring its duplication during each cell cycle as the centrioles themselves duplicate. The centrosome duplicates and splits into two equal parts during interphase, each half containing a duplicated centriole pair. These two daughter centrosomes move to opposite sides of the nucleus when mitosis begins, and they form the two poles of the mitotic spindle. A centriole consists of a short cylinder of modified microtubules, plus a large number of accessory proteins. The molecular basis for its duplication is not known. (Figure 16-30b Molecular Biology of the Cell - http://www.ncbi.nlm.nih.gov/books/NBK26809/#A3006)

Microtubule assembly and disassembly occur preferentially at the (+) end 1. Microtubules assemble by polymerization of αβ-tubulin dimers. The kinetics of tubulin polymerization and the structural intermediates observed during microtubule assembly or disassembly show that microtubule assembly is similar in many respects to microfilament assembly. ① First, at αβ-tubulin concentrations above the critical concentration (Cc), the dimers polymerize

into microtubules, while at concentrations below the Cc, microtubules depolymerize (Figure 18-8a 6th ed).

② Second, the addition of fragments of flagellar or other microtubules to a solution of αβ-tubulin accelerates the initial polymerization rate by acting as nucleation sites.

③ Third, at αβ-tubulin concentrations higher than the Cc for polymerization, dimers add to both ends of a growing microtubule, but the addition of tubulin subunits occurs preferentially at one end.

2. What determines whether the ends grow or shrink is entirely dependent on the cytosolic concentration of available monomer subunits in the surrounding area. Both the plus and the minus ends have a different critical concentration (CC). Examples in which the cytosolic concentration can affect the critical concentrations are as followed: A cytosolic concentration of subunits above both the CC+ and CC− ends results in subunit addition at both ends. A cytosolic concentration of subunits below both the CC+ and CC− ends results in subunit removal at both ends Both the plus and minus ends have different CC values and generally, the plus end will always have a lower CC value than the minus end. This is due to the increased ease of subunit addition to the plus end, leading to faster growth. Note that the cytosolic concentration of the monomer subunit between the CC+ and CC− ends is what is defined as treadmilling in which there is growth at the plus end, and shrinking on the minus end (Figure 18-8b,c 6th ed, Wikipedia).

Experimental Figure 18.10 Fluorescence microscopy reveals growth and shrinkage of individual microtubules in vivo.

Figure 18.9 Dynamic instability of microtubules in vitro.

Dynamic instability of microtubules in vitro Individual microtubule behavior was examined in a very simple experiment. Microtubules were assembled in vitro and then sheared to break them into shorter pieces whose individual lengths could be analyzed by microscopy. Under these conditions, one would expect all the short microtubules to either grow or shrink, depending on the free tubulin concentration. However, the investigators found that some of the microtubules grew in length, whereas others shortened very rapidly-thus indicating the existence of two distinct populations of microtubules. Further studies showed that individual microtubules could grow and then suddenly experience a catastrophe to a shrinking phase during which the microtubule undergoes rapid depolymerization. Moreover, sometimes a depolymerizing microtubule end could go through a rescue and begin growing again (Figure 18-9). Although this phenomenon was first seen in vitro, analysis of fluorescently labeled tubulin microinjected into live cells showed that microtubules in cells also undergo periods of growth and shrinkage (Figure 18-10). This process of alternating between growing and shrinking states is known as dynamic instability. Thus the dynamic life of a microtubule end is determined by the rate of growth, the frequency of catastrophes, the rate of depolymerization, and the frequency of rescues. As we see later, these features of microtubule dynamics are controlled in vivo. Since the (-) ends of microtubules in animal cells are generally anchored on an MTOC, this dynamic nature is most relevant to the (+) end of the microtubule.

Figure 18.11 Dynamic instability depends on the presence or absence of a GTP-β-tubulin cap.

Dynamic instability depends on the presence or absence of a GTP-β-tubulin cap A growing microtubule has a relatively blunt end, whereas a depolymerizing end has protofilaments peeling off like rams’ horns (Figure 18-11). In fact, the growing microtubule end is not simply a blunt end but rather a short and flat sheet-like structure, made by the addition of tubulin dimers to the ends of protofilaments, that then rolls up along the seam to make the cylindrical microtubule. Recent studies have provided a simple structural explanation for the two classes of microtubule ends. As we noted above, the β-subunit of the αβ-tubulin dimer is exposed on the (+) end of each protofilament. Using a GDP analog, researchers have found that artificially made single protofilaments-where there are no lateral interactions-made up of repeating αβ-tubulin dimers containing GDP-β-tubulin are straight. Thus growing microtubules with blunter ends terminate in GTP-β-tubulin, whereas shrinking ones with curled ends terminate in GDP-β-tubulin. Therefore, if the GTP molecules in the terminal β-tubulins become hydrolyzed on a microtubule will curl and a catastrophe ensues. These relationships are summarized in Figure 18-11. These results have an additional implication, and to understand this we have to consider the growing microtubule in more detail. The addition of a dimer to the (+) end of a protofilament on a growing microtubule involves an interaction between the preexisting terminal β-subunit and the new α-subunit. This interaction enhances the hydrolysis of the GTP to GDP in the formerly terminal β-subunit. However, the β-tubulin in the newly added dimer contains GTP. Thus each protofilament in a growing microtubule has mostly GDP-β-tubulin down its length and is capped by one or two terminal dimers containing GTP-β-tubulin. As we mentioned above, an isolated protofilament containing GDP-β-tubulin is curved along its length, so when it is present in a microtubule, why doesn’t‘ it break out and peel away? The lateral protofilament-protofilament interactions in the β-tubulin-GTP cap are sufficiently strong that they do not allow the microtubule to unpeel at its end-end so the protofilaments behind the GTP-β-tubulin cap are constrained from unpeeling (Figure 18-11). The energy released by GTP hydrolysis of the subunits behind the cap is stored within the lattice as structural strain waiting to be released when the GTP-β-tubulin cap is lost. If the GTP-β-tubulin cap is lost, the stored energy can do work if some structure, such as a chromosome, is attached to the disassembling microtubule end. As we will see, this stored energy contributes to the movement of chromosomes during the anaphase stage of mitosis

Figure 18.15 Proteins that destabilize the ends of microtubules.

Proteins that destabilize the ends of microtubules Various mechanisms for microtubule destabilization are known. 1. One of these involves the kinesin-13 family of proteins. Most kinesins are molecular motors, but the

kinesin-13 proteins are a distinct class that bind and curve the end of the tubulin protofilaments into the GDP-β-tubulin conformation. They then facilitate the removal of terminal tubulin dimers, therby greatly enhancing the frequency of catastrophes (Figure 18-15a). They act catalytically in the sense that they need to hydrolyze ATP to sequentially remove terminal tubulin dimers.

2. Another protein, known as Op18/stathmin, also enhances the rate of catastrophes. It was originally identified as a protein highly overexpressed in certain cancers; hence part of its name (Oncoprotein 18). Op18/stathmin is a small protein that binds two tubulin dimers in a curved, GDP-β-tubulin-like conformation (Figure 18-15b). It may function by enhancing the hydrolysis of the GTP in the terminal tubulin dimer and aiding in its dissociation form the end of the microtubule. As might be expected for a regulator of microtubule ends, it is subject to negative regulation by phosphorylation by a large variety of kinases. In fact, it has been found that Op18/stathmin is inactivated by phosphorylation near the leading edge of motile cells, which contributes to preferential growth of microtubules toward the front of the cell.

Figure 18.2 Microtubules are found in many different locations, and all have similar structures.

Experimental Figure 18.16 The rate of axonal transport in vivo can be determined by radiolabeling and gel electrophoresis.

Experimental Figure 18.17 DIC microscopy demonstrates microtubule-based vesicle transport in vitro.

Axonal transport along microtubules 1. In the early days of electron microscopy, cell biologists noted long tubules in the cytoplasm that they

called microtubules. Morphologically similar microtubules were seen making up the fibers of the mitotic spindle, as components of axons, and as the structural elements in cilia and flagella (Figure 18-2a, b). A careful examination of single microtubules from various sources seen in transverse section indicated that they are all made up of 13 longitudinal repeating units (Figure 18-2c), now called protofilaments, suggesting the various microtubules all have a common structure.

2. The results of classic pulse-chase experiments, in which radioactive precursors were microinjected into the dorsal-root ganglia near the spinal cord and then tracked along their nerve axons, showed that axonal transport occurs from the cell body down the axon. Other experiments showed transport can also occur in the reverse direction, i.e., toward the cell body. Anterograde transport proceeds form the cell body to the synaptic terminals and is associated with axonal growth and the delivery of synaptic vesicles. In the opposite, retrograde, direction, “old” membranes from the synaptic terminals move along the axon rapidly toward the cell body, where they may be degraded in lysosomes. Finding from such experiments also revealed that different materials move at different speeds (Figure 18-16).

3. Neurobiologists have long made extensive use of the squid giant axon for studying organelle movement along microtubules. Involved in regulating the squid’s water propulsion system, the aptly named giant axon can be up to 1 mm in diameter, which is about 100 times wider than the average mammalian axon. Moreover, squeezing the axon like a tube of toothpaste results in the extrusion of the cytoplasm (also known as axoplasm), which can then be observed by video microscopy. The movement of vesicles along microtubules in this cell-free system requires ATP, its rate is similar to that of fast axonal transport in intact cells, and it can proceed in both the anterograde and the retrograde directions (Figure 18-17a). Electron microscopy of the same region of the axon cytoplasm reveals organelles attached to individual microtubules (Figure 18-17b). These pioneering in vitro experiments extablished definitively that organelles move along individual microtubules and that their movement requires ATP.

Figure 18.27 Organelle transport by microtubule motors.

Organelle transport by microtubule motors Both dynein and kinesin family members play important roles in the microtubule-dependent organization of organelles in cells (Figure 18-27). Because the orientation of microtubules is fixed by the MTOC, the direction of transport-toward or away from the cell center-depends on the motor protein. 1. For example, the Golgi apparatus collects in the vicinity of the centrosome, where the (-) ends of

microtubules lie, and is driven there by dynein-dynactin. In addition, secretory cargo emerging from the endoplasmic reticulum is transported to the Golgi by dynein-dynactin. Some organelles of the endocytic pathway are associated with dynein-dynactin, including late endosomes and lysosomes.

2. Conversely, the endoplasmic reticulum is spread throughout the cytoplasm and is transported there by kinesin-1, which moves toward the peripheral (+) ends of microtubules. Kinesins have been shown to transport mitochondria, as well as nonmembranous cargo such as specific mRNAs encoding proteins that need to be localized during development.

Figure 18.19 Model of kinesin-1-catalyzed vesicle transport.

Figure 18.18 Structure of kinesin-1.

Model of kinesin-1-catalyzed vesicle transport 1. Kinesin-1-dependent movement of vesicles can be tracked by in vitro motility assays similar to those used to study myosin-dependent movements. In one type of assay, a vesicle or a plastic bead coated with kinesin-1 is added to a glass slide along with a preparation of stabilized microtubules. In the presence of ATP, the beads can be observed microscopically to move along a microtubule in one direction. Researchers found that the beads coated with kinesin-1 always moved from the (-) to the (+) end of a microtubule (Figure 18-19). Thus kinesin-1 is a (+) end-directed microtubule motor protein, and additional evidence shows that it mediates anterograde axonal transport. 2. Kinesin-1 isolated from squid giant axons is a dimer of two heavy chains, each associated with a light chain, with a total molecular weight of about 380,000. The molecule comprises a pair of globular head domains connected by a short flexible linker domain to a long central stalk and terminating in pair of small globular tail domains, which associate with the light chains (Figure 18-18). Each domain carries out a particular function: ① the head domain binds microtubules and ATP and is responsible for the motor activity of kinesin; ② the linker domain is critical for forward motility; ③ the stalk domain is involved in dimerization of the two heavy chains; and ④ the tail domain is responsible for binding to receptors on the membrane of cargoes.

Figure 18.20 Structure and function of selected members of the kinesin superfamily.

Figure 18.21 Kinesin-1 uses ATP to “walk” down a microtubule.

Figure 18.22 Convergent structural evolution of the ATP-binding cores of myosin and kinesin heads.

Structure and function of kinesin 1. As with the different classes of myosin motors, in the various kinesin families the conserved motor domain is fused to a variety of class-specific nonmotor domains (Figure 18-20). Whereas kinesin-1 has two identical heavy chains and two identical light chains, members of the kinesin-2 family (also involved in organelle transport) have two different related heavy-chain motor domains and a third polypeptide that associates with the tail and binds cargo. Members of the bipolar kinesin-5 family have four heavy chains, forming bipolar motors that can cross-link antiparallel microtubules and, by walking toward the (+) end of each microtubule; this class functions in mitosis. Yet another type, the kinesin-13 family, has two subunits but with the conserved kinesin domain in the middle of the polypeptide. Kinesin-13 proteins do not have motor activity, but recall that these are special ATP-hydrolyzing proteins that can enhance the depolymerization of microtubule ends. 2. The ATP cycle of kinesin-1 movement is most easily understood by first considering the cycle just after the motor has taken a step (Figure 18-21a). At this point the motor has a nucleotide-free leading head, under which conditions it is strongly bound to a tubulin dimer in a protofilament, and an ADP-bound trailing head that is weakly associated with the protofilament. ① The leading head then binds ATP (step 1), ② which induces a conformational change that causes the yellow linker region to swing forward and dock

into its associated head domain, thereby thrusting the trailing head forward (step 2). ③ The new leading head now finds a binding site 16 nm down the microtubule, to which it binds weakly

(step 3). ④ The leading head now releases ADP and binds tightly to the microtubule, which induces the trailing

head to hydrolyze ATP to ADP and Pi (step 4). Pi is released and the trailing head is converted into a weak binding state, and also releases the docked linker domain. The cycle now repeats itself for another step.

3. When the x-ray structure of the kinesin head was determined, it revealed a major surprise-the catalytic core has the same overall structure as myosin’s (Figure 18-22). This occurs despite the fact that there is no amino acid sequence conservation between the two proteins, arguing strongly that convergent evolution twice generated a fold that can utilize the hydrolysis of ATP to generate work.

Lodish • Berk • Kaiser • Krieger • scott • Bretscher • Ploegh • Matsudaira

MOLECULAR CELL BIOLOGY SEVENTH EDITION

CHAPTER 18 Cell Organization and Movement II:

Microtubules and Intermediate Filaments

Copyr ight © 2013 by W. H. Freeman and Company

Figure 18.23 The domain structure of cytoplasmic dynein.

Figure 18.24 The power stroke of dynein.

Figure 18.25 The dynactin complex linking dynein to cargo.

Dynein 1. The second family of microtubule motor proteins, the dyneins, is responsible for retrograde axonal

transport, transit of Golgi vesicles to the centrosome, and some other (-) end–directed movements. They are composed of two or three heavy chains complexed with a poorly determined number of intermediate and light chains. Here we consider cytosolic dynein, which has a role in the movement of vesicles and chromosomes.

2. Like kinesin I, cytosolic dynein is a two-headed molecule, with two identical or nearly identical heavy chains forming the head domains. A single dynein heavy chain consists of a number of distinct domains (Figure 18-23). It consists of the stem, to which the other dynein subunits bind and which associates with its cargo through dynactin. The next part of the heavy chain is a linker that plays a critical role during ATP-dependent motor activity. A large part of the heavy chain makes up the head containing the AAA ATPase domain, consisting of six repeats that assemble into a flowerlike structure, within which lies the ATPase activity. Embedded between the fourth and fifth AAA repeats is the stalk, which protrude from the structure and contains the microtubule-binding region.

3. Before a power stroke, the stem is attached to the linker that lies across the AAA domain and associates with the first and third AAA repeats (Figure 18-24a and b, left panels). Upon ATP binding and hydrolysis, the AAA ring changes conformation slightly and the linker domain becomes associated with the first and fifth AAA repeat. This conformational change rotates the molecule to bring the stem and stalk closer, resulting in the transport of cargo toward the (-) end of the microtubule (Figure 18-24a and b, right panels).

4. Unlike kinesin-1, dynein cannot mediate cargo transport by itself. Rather, dynein-related transport requires dynactin, a large protein complex that links vesicles and chromosomes to the dynein light chains (Figure 18-25).

The EMBO Journal (2011) 30, 3527–3539

Subunit composition of dynein and kinesin-1 Schematic representation of the subunit composition of dynein and kinesin-1. (A) The motor containing cytoplasmic dynein heavy chains are shown in orange and associated intermediate and light chains in shades of blue. The motor domain is composed of six AAA ATPase domains arranged in a hexameric ring from which a microtubule binding stalk projects. The N-terminal tail of the heavy chain mediates its dimerization and contains the binding sites for two intermediate chains (ICs) and two light intermediate chains (LICs). The two intermediate chains (ICs) also interact with three pairs of light chains: Tctex, LC7 and LC8. (B) Kinesin-1 is a heterotetramer composed of two motor containing heavy chains (orange) and two light chains (blue). The microtubule binding motor domain is found in the N-terminus of the heavy chain. The light chains associate with the heavy chains via heptad repeat regions in their N-terminus. The C-terminal half of the light chains is composed of six tetratricopeptide repeats (TPR), which represent cargo binding domains (The EMBO Journal (2011) 30, 3527–3539).

Figure 18.28 Movement of pigment granules in frog melanophores.

Movement of pigment granules in frog melanophores Melanophores are cells of the vertebrate skin that contain hundreds of dark melanin-filled pigment granules called melanosomes. The movement of the granules is mediated by changes in intracellular cAMP and is dependent on microtubules. Studies investigating which motors are involved have shown that pigment granule dispersion requires kinesin-2, whereas aggregation requires cytoplasmic dynein-dynactin.

Experimental Figure 18.31 Video microscopy shows flagellar movements that propel sperm and Chlamydomonas forward.

Figure 18.30 Structural organization of cilia and flagella.

Figure 18.32 Cilia and flagella bending mediated by axonemal dynein.

Cilia and flagella bending mediated by axonemal dynein 1. Ciliary and flagellar beating is characterized by a series of bends, originating at the base of the structure and propagated toward the tip (Figure 18-31). The bends push against the surrounding fluid, propelling the cell forward or moving the fluid across a fixed epithelium. A bend results from the sliding of adjacent doublet microtubules past one another. 2. Cilia and flagella are flexible membrane extensions that project from certain cells. They range in length from a few micrometers to more than 2 mm for some insect sperm flagella. Virtually all eukaryotic cilia and flagella possess a central bundle of microtubules, called the axoneme, which consists of nine doublet microtubules surrounding a central pair of singlet microtubules (Figure 18-30). This characteristic “9 + 2” arrangement of microtubules is seen in cross section with the electron microscope. Each doublet microtubule consists of A and B tubules. The (+) end of axonemal microtubules is at the distal end of the axoneme. At its point of attachment to the cell, the axoneme connects with the basal body. Containing nine triplet microtubules, the basal body plays an important role in initiating the growth of the axoneme. The axoneme is held together by three sets of protein cross-links (Figure 18-30a). The central pair of singlet microtubules is connected by periodic bridges, like rungs on a ladder, and is surrounded by a fibrous structure termed the inner sheath. A second set of linkers, composed of the protein nexin, joins adjacent outer doublet microtubules. Radial spokes, which radiate from the central singlets to each A tubule of the outer doublets, are proposed to regulate dynein. Permanently attached periodically along the length of the A tubule of each doublet microtubule are inner-arm and outer-arm dyneins (Figure 18-30a). 3. A clue to how this occurs came from studies of isolated axonemes. In classic experiments, axonemes were gently treated with a protease that cleaves just the nexin links. When ATP was added to the treated axonemes, the doublet microtubules slid past one another as dynein, attached to the A tubule of one doublet, “walked” down the B tubule of the adjacent doublet (Figure 18-32b, c). In an axoneme with intact nexin links, the action of dynein induces flagellar bending as the microtubule doublets are connected to one another (Figure 18-32a).

Figure 18.33 Intraflagellar transport.

Figure 18.34 Many interphase cells contain a non-motile primary cilium.

Cilia and flagella 1. Cilia and flagella have a mechanism, intraflagellar transport (IFT), to transport material to their tips by

kinesin-2 and from the tip back to the base by cytoplasmic dynein. This transport regulates the function and length of cilia and flagella (Figure 18-33).

2. Many cells have on their surface a single non-motile primary cilium that lacks the normal central pair of microtubules and dynein side arms of motile cilia. The primary cilium functions as a sensory organelle, with receptors for extracellular signals localized to its plasma membrane. Due to its sensory function, many diseases result from defects in receptor localization or in the structure of the primary cilium itself (Figure 18-34).

Figure 19.2 The stages of mitosis.

Figure 18.36 The stages of mitosis.

Figure 18.35 Relation of centrosome duplication to the cell cycle.

The stages of mitosis 1. Figure 18-36 depicts the characteristic series of events that can be observed by light microscopy in

mitosis in a eukaryotic cell. Although the events unfold continuously, they are conventionally divided into four substages: prophase, metaphase, anaphase, and telophase (Figure 19-2). The beginning of mitosis is signaled by the appearance of condensing chromosomes, first visible as thin threads inside the nucleus. By late prophase, each chromosome appears as two identical filaments, the chromatids (often called sister chromatids), held together at a constricted region, the centromere. Each chromatid contains one of the two new daughter DNA molecules produced in the preceding S phase of the cell cycle; thus each cell that enters mitosis has four copies of each chromosomal DNA, designated 4n.

2. Mitosis is the process that partitions newly replicated chromosomes equally into separate parts of a cell. The last step in the cell cycle, mitosis takes about 1 hour in an actively dividing animal cell. In that period, the cell builds and then disassembles a specialized microtubule structure, the mitotic apparatus. Larger than the nucleus, the mitotic apparatus is designed to attach and capture chromosomes, align the chromosomes, and then separate them so that the genetic material is evenly partitioned to each daughter cell. The structure of the mitotic apparatus changes constantly during the course of mitosis. Because each half of the metaphase mitotic apparatus emanates from a polar centrosome, its assembly depends on duplication of the centrosome and movement of the daughter centrosomes to opposite halves of the cell. This process, known as the centriole cycle (or centrosome cycle) marks the first steps in mitosis, beginning during G1 when the centrioles and other centrosome components are duplicated (Figure 18-35). By G2, the two “daughter” centrioles have reached full length, but the duplicated centrioles are still present within a single centrosome. Early in mitosis, the two pairs of centrioles separate and migrate to opposite sides of the nucleus, establishing the bipolarity of the dividing cell.

Figure 18.37 Mitotic spindles have three distinct classes of microtubules.

Mitotic spindles have three distinct classes of microtubules At metaphase, the mitotic apparatus is organized into two parts: a central mitotic spindle and a pair of asters (Figure 18-37a). The spindle is a bilaterally symmetric bundle of microtubules and associated proteins with the overall shape of a football; it is divided into opposing halves at the equator of the cell by the metaphase chromosomes. An aster is a radial array of microtubules at each pole of the spindle. In each half of the spindle, a single centrosome at the pole organizes three distinct sets of microtubules whose (-) ends all point toward the centrosome (Figure 18-37b).

1. One set, the astral microtubules, forms the aster; they radiate outward from the centrosome toward the

cortex of the cell, where they help position the mitotic apparatus and later help to determine the cleavage plane in cytokinesis. The other two sets of microtubules compose the spindle.

2. The kinetochore microtubules attach to chromosomes at specialized attachment sites on the chromosomes called kinetochores.

3. Polar microtubules do not interact with chromosomes but instead overlap with polar microtubules from the opposite pole. Two types of interactions hold the spindle halves together to form the bilaterally symmetric mitotic apparatus: (1) lateral interactions between the overlapping (+) ends of the polar microtubules and (2) end-on interactions between the kinetochore microtubules and the kinetochores of the sister chromatids. The large protein complexes, called cohesins, that link sister chromatids together are discussed.

Figure 18.39 The structure of a mammalian kinetochore.

The structure of a mammalian kinetochore The linkage of metaphase chromosomes to the (+) ends of kinetochore microtubules is mediated by a large protein complex, the kinetochore, which has several functions: to trap and attach microtubule ends to the chromosomes, to generate force to move chromosomes along microtubules, and to regulate chromosome separation and translocation to the poles. In an animal cell, the kinetochore forms at the centromere and is organized into an inner and outer layer embedded within a fibrous corona (Figure 18-39). In all eukaryotes, three components participate in attaching chromosomes to microtubules: the centromere, kinetochore and spindle proteins, and the cell-cycle machinery. The location of the centromere and hence that of the kinetochore is directly controlled by a specific sequence of chromosomal DNA termed centromeric DNA.

Figure 18.40 Chromosome capture and congression in prometaphase.

Chromosome capture and congression in prometaphase 1. How does a kinetochore become attached to microtubules in prometaphase? ① Microtubules nucleated from the spindle poles are very dynamic, and when they contact the

kinetochore, either laterally or at their end, this can lead to chromosomal attachment (Figure 18-40a, step 1a and 1b). Microtubules “captured” by kinetochores are selectively stabilized by reducing the level of catastrophes, thereby promoting the chance that the attachment will persist.

② Once a kinetochore is attached laterally or terminally to a microtubule, the motor protein dynein-dynactin associates with the kinetochore to move the duplicated chromosome down the microtubule toward the spindle pole. This eventually results in an end-on attachment of the microtubule to one kinetochore (Figure 18-40a, step 2).

③ This movement helps orient the sister chromatid so that the unoccupied kinetochore on the opposite side is pointing toward the distal spindle pole. Eventually a microtubule from the distal pole will capture the free kinetochore; at this point the sister chromatid pair is now said to be bi-oriented (Figure 18-40a, step 3).

④ With the two kinetochores attached to opposite poles, the duplicated chromosome is now under tension, being pulled in both directions by the two sets of kinetochore microtubules. When one or a few chromosomes are bi-oriented, other chromosomes use these existing kinetochore microtubules to contribute to their orientation and movement to the spindle center. This is mediated by kinesin-7 associated with the free kinetochore moving the chromosome to the (+) end of the kinetochore microtubule (Figure 18-40a, step 4).

2. During prometaphase, the chromosomes come to lie at the midpoint between the two spindle poles, in a process known as chromosome congression. During this process, bi-oriented chromosome pairs often oscillate backward and forward before arriving at the midpoint. Chromosome congression involves the coordinated activity of several microtubule-based motors together with regulators of microtubule assembly and disassembly (Figure 18-40b). On the shortening side, a kinesin-13 protein stimulates disassembly and a dynein-dynactin complex moves the chromosome toward the pole. On the side with lengthening microtubules, kinesin-7 protein holds on to the growing microtubule. Kinesin-4, a (+) end-directed motor, interacts with the polar microtubules to pull the chromosomes toward the center of the spindle.

Figure 18.41 CPC regulation of microtubule-kinetochore attachment.

CPC regulation of microtubule-kinetochore attachment CPC regulation of microtubule-kinetochore attachment. The chromosomal passenger complex (CPC) associated with the inner kinetochore plate keeps microtubule attachments weak by the activity of its kinase component Aurora B, which phosphorylates critical kinetochore proteins. When a chromosome is bi-oriented, tension is generated and the Aurora B substrates are pulled away from the kinase (Figure 18-41). Without phosphorylation of kinetochore proteins by Aurora B, the chromosome-kinetochore attachment becomes stable.

Figure 18.42 Chromosome movement and spindle pole separation in anaphase.

Chromosome movement and spindle pole separation in anaphase The same forces that form the spindle during prophase and metaphase also direct the separation of chromosomes toward opposite poles at anaphase. Anaphase is divided into two distinct stages, anaphase A and anaphase B (early and late anaphase). Anaphase A is characterized by the shortening of kinetochore microtubules at their (+) ends, which pulls the chromosomes toward the poles. In anaphase B, the two poles move farther apart, bringing the attached chromosomes with them into what will become the two daughter cells. 1. Microtubule shortening in anaphase A. In one such study, purified microtubules were mixed with

purified anaphase chromosomes; as expected, the kinetochores bound preferentially to the (+) ends of the microtubules. To induce depolymerization of the microtubules, the reaction mixture was diluted, thus lowering the concentration of free tubulin dimers. Video microscopy analysis then showed that the chromosomes moved toward the (-) end, at a rate similar to that of chromosome movement during anaphase in intact cells. shortening of kinetochore microtubules at their (+) ends moves chromosomes toward the poles in mammalian cells. One of the kinesin-13 proteins is localized at the kinetochore and enhances disassembly there (Figure 18-42, A1), and the other is localized at the spindle pole, enhancing depolymerization there (Figure 18-42, A2). Thus, anaphase A is powered in part by kinesin-13 proteins specifically localized at the kinetochore and spindle pole to shorten the kinetochore microtubules at both their (+) and (-) ends, drawing the chromosomes to the poles.

2. The second part of anaphase involves separation of the spindle poles in a process known as anaphase B. A major contributor to this movement is the involvement of the bipolar kinesin-5 proteins (Figure 18-42, B1). These motors associate with the overlapping polar microtubules, and since they are (+) end-directed motors, they push the poles apart. While this is happening, the polar microtubules have to grow to accommodate the increased distance between the spindle poles. Another motor-the microtubule (-) end-directed motor cytoplasmic dynein, localized and anchored on the cell cortex-pulls on the astral microtubules and thus helps separate the spindle poles (Figure 18-42, B2).