Microtubules and Centrosomes

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CHAPTER 34 Microtubules and Centrosomes

Microtubules are stiff, cylindrical polymers of α- and β-tubulin (Fig. 34-1) that provide support for a variety of cellular components and tracks for movements powered by motor proteins called kinesins and dyneins. Microtubules are 25nm in diameter and can grow longer than 20μm in cells and 3mm in vitro. The head-to-tail arrangement of dimers of α- and β-tubulin in the wall of microtubules give the polymer a molecular polarity. The “plus” (β-tubulin) end grows faster than the “minus” end.

Figure 34-1 microtubules (arrows) visualized in electron micrographs of thin sections of a cultured mammalian cell. A, Longitudinal section.B, Cross section.Insets, Electron micrographs of microtubules in frozen hydrated rat hepatoma cells.

(Main panels, Courtesy of R. Goldman, Northwestern University, Evanston, Illinois. Insets, Courtesy of Cédric Bouchet-Marquis and Jacques Dubochet, University of Lausanne, Switzerland.)

A great simplifying principle is that microtubules have a radial organization in many types of cells (Fig. 34-2A). Typically, the plus end is peripheral, and the minus end is anchored in a microtubule-organizing center. One exception to this radial organization is found in dendrites of nerve cells, where about 40% of the microtubules are oriented with the minus end away from the cell body.

Figure 34-2 arrangements of microtubules in various cells. a–c, Fluorescence micrographs of microtubules stained with antibodies to tubulin.A, Vertebrate tissue culture cells.Green microtubules radiate from thered centrosome near the blue nucleus of an interphase cell (upper panel). A HeLa cell in mitosis with green microtubules radiating from the two poles toward the blue chromosomes and red centromeres (stained with anticentromere antibody) (lower panel). B, Columnar epithelial cells in tissue culture. Three-dimensional reconstructions show microtubules oriented along the long axis of the cell.C, Plant cells. Maize epidermal cells with green microtubules in the cortex and in the mitotic spindles of three dividing cells near the middle of the field. Nuclei are stainedorange. D, Live budding yeast. Two interphase cells with microtubules radiating from the spindle pole bodies (arrows) associated with the nuclear envelope; these microtubules are marked with dynein fused to green fluorescent protein (upper panel). A cell late in mitosis with a bundle of microtubules extending from one spindle pole body to the other inside the nucleus; these microtubules are marked with tubulin fused to green fluorescent protein (lower panel).

(A, Upper panel, Courtesy of A. Khodjakov, Wadsworth Center, Albany, New York; lower panel, Courtesy of D. W. Cleveland, University of California, San Diego. B, Courtesy of R. Bacallao, University of Indiana Medical School, Indianapolis. C, Courtesy of L. Smith, University of California, San Diego. D, Courtesy of P. Maddox, University of North Carolina, Chapel Hill.)

In most animal cells, the organizing center for cytoplasmic microtubules is the centrosome. Centrosomes consist of centrioles and a surrounding matrix containing the active component in microtubule nucleation—a complex of proteins including the specialized tubulin isoform γ-tubulin. The microtubule-organizing center is more diffuse in columnar epithelial cells, where microtubules originate from a broad zone containing γ-tubulin near the apex of the cell (Fig. 34-2B). In plant cells, γ-tubulin and microtubules are found throughout the cortex rather than in a discrete array, although they form a bipolar mitotic spindle during cell division (Fig. 34-2C). Microtubules in fungi grow from a spindle pole body, an organizing center containing γ-tubulin that is associated with the nuclear envelope (Fig. 34-2D). Animals and protozoa with cilia and flagella use basal bodies (Fig. 34-3B) to nucleate the assembly of microtubules for their motile structure, called an axoneme.

Figure 34-3 cilia on mussel gill epithelial cells. A, A scanning electron micrograph reveals how the bending movements of the thread-like cilia are coordinated in waves.B–C, Transmission electron micrographs of cilia.B, Longitudinal section showing basal bodies and proximal parts of two cilia.C, Cross section of many cilia showing the nine outer doublets and the central pair of microtubules.

(Courtesy of P. Satir, Albert Einstein College of Medicine, Bronx, New York. Reference: Satir P: How cilia move. Sci Am 231:44–52, 1974.)

Microtubules vary considerably in stability. Those that form the axonemes in eukaryotic cilia and flagella are stable for days to weeks. Cytoplasmic microtubules turn over much more rapidly, within minutes in the case of the interphase array of microtubules and within tens of seconds for mitotic spindle microtubules. These dynamic microtubules randomly undergo rapid depolymerization and then regrow over a period of seconds to minutes. This “dynamic instability” helps to remodel the network of microtubules in cytoplasm and contributes to some forms of motility, including the assembly of the mitotic spindle and movements of chromosomes during mitosis (see Fig. 44-7).

Because the same tubulin dimers can form dynamic single microtubules in cytoplasm and stable doublet microtubules in axonemes, it is believed that accessory proteins specify both the stability and diverse structures assembled from tubulin. The various families of microtubule-associated proteins (MAPs) bind tubulin dimers, stabilize polymers, associate with microtubule ends, or sever cytoplasmic microtubules. In the axonemes of cilia and flagella, more than 100 accessory proteins organize and stabilize the regular array of nine outer doublet microtubules and two central single microtubules (see Fig. 38-16).

Microtubule motor proteins (see Figs. 36-13 and 36-14) power movements ranging from the slow movements of chromosomes on the mitotic spindle (see Fig. 44-14) to the rapid beating of cilia and flagella (see Fig. 38-14). Different motors move toward the plus and minus ends of microtubules. The main minus-end-directed motor, dynein, drives the beating of cilia and flagella. Dynein and the kinesin family of plus-end motors move membrane-bound organelles, RNA particles, viruses, and other cargo along microtubules (see Fig. 37-7). These active movements determine, to a great extent, the distribution of cellular organelles and the shape of cells.

Tubulin Structure

The tubulin molecule is a heterodimer of a and β subunits that share a common fold and 40% identical residues (Fig. 34-4). Dimers of α- and β-tubulin are stable and rarely dissociate at the 10- to 20-mM concentr-tions of tubulin found in cells. γ-tubulins have the same fold.

Figure 34-4 structure of the α-tubulin/β-tubulin dimer. A, Ribbon diagrams with space-filling GTP on α-tubulin and GDP on β-tubulin.B, Surface rendering. This historic structure was the first high-resolution structure of a nonmembrane protein to be determined by electron crystallography, using sheets of tubulin protofilaments as the specimen. Each subunit consists of about 450 residues arranged in two domains. Each domain is a β-sheet flanked by a-helices. The nucleotides bind in pockets similar to the binding site for nicotinamide-adenine dinucleotide (NAD) on the enzyme glyceraldehyde-3-phosphate dehydrogenase.

(PDB file: 1TUB. Reference: Nogales E, Downing K: Structure of the a/b tubulin dimer by electron crystallography. Nature 391:199–203, 1998.)

Each tubulin subunit binds a guanine nucleotide, either guanosine triphosphate (GTP) or guanosine diphosphate (GDP). The fold and the GTP-binding site of tubulin do not resemble those of other GTP-binding proteins (see Fig. 25-7). GTP on α-tubulin is buried in the dimer, so it does not exchange with solution GTP; hence, it is called the nonexchangeable N-site. GTP on β-tubulin is exposed in the dimer and exchanges slowly (K_d = 50nM), so this is known as the exchangeable site, or E-site. When incorporated into a microtubule, contacts with the adjacent a subunit bury the GTP on the β subunit and promote its hydrolysis. Neither bound guanine nucleotide can exchange when tubulin is buried in the wall of a microtubule. As is explained later in this chapter, the nature of the nucleotide on the β subunit profoundly affects microtubule assembly.

Most Archaea and Bacteria have a protein called FtsZ with the same fold as tubulin. The two proteins are most likely to have had a common ancestor, but their sequences have diverged considerably. FtsZ also forms polymers and is required for cytokinesis of prokaryotes (see Fig. 44-21). One group of Bacteria lost their FtsZ but have acquired genes for both α- and β-tubulin, most likely by lateral transfer from an eukaryote.

Tubulins are modified in a number of different ways in cells. Over time, stable microtubules accumulate two modifications—acetylation of lysine-40 and removal of the C-terminal tyrosine of α-tubulin—but neither modification is responsible for the stability. The enzyme carboxypeptidase B removes the tyrosine, leaving a glutamic acid exposed. Another enzyme, tyrosine-tubulin ligase, can replace the tyrosine. Other microtubules are modified by the addition of a polymer of up to six glutamic acid residues to the γ-carboxyl groups of glutamic acid residues of both α-tubulin and β-tubulin. Addition of one or more glycines to the γ-carboxyl of other glutamate residues stabilizes the central pair of microtubules in axonemes (see Fig. 38-16).

Tubulin Diversity

Tubulins are remarkably conserved across the phylogenetic tree; more than 75% of the residues of animal α- or β-tubulins are identical to their plant homologs. On the other hand, species differ considerably in their variety of tubulin isoforms. Vertebrates have six to eight genes each encoding variants of α- and β-tubulin, whereas budding yeast have but two α-tubulin genes and one β-tubulin gene. Unicellular ciliates such as Tetrahymena assemble a greater variety of microtubule-based structures than humans have in their various tissues, but they do this using only one α- and one β-tubulin polypeptide. Most vertebrate cells express several tubulin isoforms, but exceptional cases, such as bird red blood cells, express a single α-tubulin and β-tubulin. It is impressive that the sequences of orthologous tubulins vary little, if at all, between different vertebrate species, whereas the paralogous β-tubulin isoforms in one organism differ by about 10% in primary structure. The sequences of γ-tubulins are also conserved between species.

Two views rationalize the significance of multiple α- and β-tubulin isoforms. On one hand, isoforms have different assembly properties and affinities for microtubule-associated proteins (MAPs) that may confer some advantage to the organism. This is illustrated by the failure of paralogous tubulin genes to substitute for each other in flies. Alternatively, the proteins themselves may be largely interchangeable (and all isoforms appear to copolymerize), but different genes may be required to ensure precise control of biosynthesis in particular cells at appropriate times during development. For example, the two α-tubulin proteins of the filamentous fungusAspergillus can substitute for each other, but two genes are required to control the expression of tubulin at specific times in the life cycle.

Four tubulin isoforms called δ-, σ-, ζ-, and η-tubulin are found in protozoa, algae, and vertebrates but are conspicuously missing from fungi and most plants. All of these isoforms are required for the structure or function of centrioles or basal bodies. Their absence accounts for the lack of centrioles in plants and fungi.

Structure of Microtubules

Microtubules are cylinders constructed of longitudin-ally oriented protofilaments with a 4-nm longitudinal repeat arising from the tubulin subunits (Fig. 34-5). Most cytoplasmic microtubules have 13 protofilaments, but microtubules in some cells have 11, 15, or 16 protofilaments. Microtubules assembled in vitro can have 11 to 15 protofilaments, but 13 is the favored number. For years, microtubules were thought to be true helices, but microtubules with 13 protofilaments have a longitudinal seam or discontinuity between two of the protofilaments, breaking up perfect helical packing of the subunits (Fig. 34-5D). The other protofilaments are aligned identically with respect to their neighbors.

Figure 34-5 microtubule structure. A, Electron micrograph of negatively stained microtubules.B–C, Longitudinal and cross sections of a reconstruction of a 13-protofilament microtubule made by image processing of electron micrographs.B, Atomic model made by fitting the tubulin dimer model into the reconstruction of the microtubule.C, Surface rendering of the reconstruction.D, Drawing of the microtubule used throughout this book. It shows the longitudinal seam between two protofilaments, which breaks the helical repeat of tubulin dimers.

(A, Courtesy of D. B. Murphy, Johns Hopkins Medical School, Baltimore, Maryland. B–C, Courtesy of R. Milligan, Scripps Research Institute, La Jolla, California. Note that a higher resolution is now available: Li H, DeRosier D, Nicholson WV et al: Microtubule structure at 8Å resolution. Structure 10:1317–1328, 2002.)

Microtubules are polar because all of the dimers have the same orientation. β-tubulin is oriented toward the plus end; α-tubulin is oriented toward the minus end. This polarity results in different rates of growth at the two ends. The plus end grows faster than the minus end. For the purpose of marking polarity in electron micrographs, microtubules in extracted cells can be decorated with either exogenous dynein or with excess tubulin, which can form curved “hooks” under nonphysiological conditions. These hooks provided the primary evidence that the minus ends are usually associated with microtubule-organizing centers.

As might be expected from their cylindrical structure, microtubules are much stiffer than are either actin filaments or intermediate filaments. If enlarged 1 million–fold to a diameter of 25μm, microtubules would have mechanical properties similar to those of a plastic pipe: quite stiff locally but flexible over distances of several meters (micrometers in the cell). On the same scale, actin filaments would be like 8-mm plastic rods, and intermediate filaments would be akin to 10-mm braided plastic ropes. Thus, microtubules are resistant to compression and support asymmetrical structures much more effectively than the other cytoskeletal polymers. Interactions of microtubules with both actin filaments and intermediate filaments reinforce the cytoskeleton.

The stiffness, length, and polarity of microtubules make them valuable both for cytoskeletal support and as tracks for microtubule-based motors. Because they resist compression, microtubules are called on more frequently than actin filaments or intermediate filaments to support asymmetrical cellular structures, including axonemes, the mitotic spindle, and elaborate surface processes of some protozoa (see Fig. 38-4). Plants provide a spectacular example of the influence of microtubules on morphology: Point mutations in tubulin influence whether climbing plants wrap in a left-handed or right-handed helix around their supports.

Microtubule Assembly from GTP Tubulin

Microtubules assemble from pure GTP-tubulin subunits much like actin filaments do using subunits with bound ATP (see Figs. 5-6 and 33-8). Superficially, the assembly of microtubules is a simple bimolecular reaction of tubulin dimers with the ends of the polymer. Association and dissociation of tubulin occurs only at the ends, not from the walls of microtubules. Growth is faster at the plus end than at the minus end (Fig. 34-6), and assembly of GTP-tubulin is much more favorable than that of GDP-tubulin. The rate of elongation is proportional to the concentration of GTP-tubulin dimers above the critical concentration at each end. The rate constants measured from the slopes and intercepts in elongation experiments (Table 34-1) are similar to those for actin (see Fig. 33-8), although this similarity might be misleading, as the number of sites growing at the end of a microtubule is unknown and is most likely to be greater than one as on actin filaments.

Figure 34-6 elongation of pure tubulin microtubules. The rate of elongation at plus and minus ends depends on the concentration of GTP-tubulin dimers. Slopes give association rate constants, x-intercepts give the critical concentrations, and the y-intercepts (after linear extrapolation[dashed lines] from positive growth rates to the y-axis) give the dissociation rate constants for GTP tubulin. The dissociation rate of GDP tubulin is shown as a single point at 733s⁻¹ on the y-axis.

(Data from Walker RA, O’Brien ET, Pryer NK, et al: Dynamic instability of individual microtubules. J Cell Biol 107:1437–1448, 1988.)

Table 34-1 RATE CONSTANTS FOR THE ASSEMBLY OF MICROTUBULES IN VITRO AND IN CELLS

Reaction	Plus End	Minus End
In Vitro Elongation of Purified Tubulin
Association of GTP tubulin	9 μM⁻¹s⁻¹	4 M⁻¹s⁻¹
Dissociation of GTP tubulin	44 s⁻¹	23 s⁻¹
Association of GDP tubulin	Unknown	Unknown
Dissociation of GDP tubulin	733 s⁻¹	915 s⁻¹
Steady-State Dynamic Instability
Frequency of catastrophe in vitro at 7 μM tubulin	0.0045 s⁻¹	0.003 s⁻¹
Frequency of rescue in vitro at 7 μM tubulin	0.02 s⁻¹	0.06 s⁻¹
Microtubule Dynamic Instability in Live Cells
Frequency of catastrophe in vivo (interphase)	0.014 s⁻¹
Frequency of catastrophe in vivo (mitosis)	0.017 s⁻¹
Frequency of rescue in vivo (interphase)	0.046 s⁻¹
Frequency of rescue in vivo (mitosis)	0

Data from Walker RA, O’Brien ET, Pryer NK, et al: Dynamic instability of individual microtubules. J Cell Biol 107:1437–1448, 1988.

Drugs (Box 34-1) and environmental conditions can depolymerize microtubules. Pioneering light microscopic observations of live cells established that mitotic spindle fibers (later shown to be microtubules) are sensitive to both cold and high hydrostatic pressure. This makes it possible to purify microtubules and tightly associated proteins by cycles of depolymerization in the cold and repolymerization at higher temperatures. Dissociated microtubule components in ice-cold cellular extracts polymerize when rewarmed to body temperature. Pelleting in a centrifuge separates microtubules and any associated proteins from soluble components. Resuspension in cold buffer with GTP depolymerizes the microtubules. The cycle is repeated until the desired degree of purity is achieved. Adaptations of tubulins or accessory proteins allow cold-water organisms to assemble microtubules at temperatures near freezing.

BOX 34-1 Pharmacologic Tools for Studying Tubulin and Microtubules

Tubulin binds several therapeutically active plant alkaloids and synthetic chemicals, including two of the most successful drugs used to treat cancer. Vinblastine (from periwinkle) interferes with microtubule dynamics by binding between tubulin dimers at the ends of microtubules. Taxol (from the bark of the Western yew) binds β-tubulin and stabilizes microtubules. At substoichiometric concentrations, vinblastine and taxol are effective in cancer chemotherapy because they interfere with the dynamic instability of mitotic spindle microtubules and block cell division. Colchicine (from the autumn crocus) and nocodazole (a synthetic chemical) inhibit microtubule assembly by binding dissociated tubulin dimers. Colchicine binds dimers between the two subunits, stabilizing a bent conformation that cannot fit into the microtubule lattice. Colchicine is used empirically to treat gout, a painful condition that results from the accumulation of uric acid crystals in joints and other tissues, but no one knows precisely how it works or why it is not more toxic.

Microtubule-Organizing Centers

Spontaneous nucleation of microtubules from tubulin dimers is so unfavorable that it is probably irrelevant to living cells. In vitro, this slow nucleation of pure tubulin accounts for a substantial lag of minutes during the formation of microtubules. Presumably, tubulin dimers form oligomers that assemble into small sheets of protofilaments and eventually close to form a cylinder, but the details are still under investigation.

Instead of spontaneous nucleation, virtually all cellular microtubules arise from microtubule-organizing centers. Microtubules in cilia and flagella grow directly from basal bodies (see Fig. 38-17), but other microtubules originate in the pericentrosomal material surrounding centrioles in the centrosome (Figs. 34-2A and 34-18), from fungal spindle pole bodies (Fig. 34-19) or nuclear envelopes and less-organized material in the cortex of some epithelial cells (Fig. 34-2B) and plant cells (Fig. 34-2C). As is described in the section on centrosomes later in this chapter, these organizing centers use assemblies of γ-tubulin to initiate microtubules.