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Alberts B, Bray D, Lewis J, et al. Molecular Biology of the Cell. 3rd edition. New York: Garland Science; 1994.

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Molecular Biology of the Cell. 3rd edition.

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Microtubules 13


Microtubules, as we have seen, are long, stiff polymers that extend throughout the cytoplasm and govern the location of membrane-bounded organelles and other cell components. In this section we discuss the assembly of these remarkable structures from tubulin molecules and explain how their polymerization and depolymerization are controlled by the nucleotide GTP. We then examine some ways in which selected microtubules are stabilized in the cell by their association with specific accessory proteins. Finally, we discuss the importance of microtubule-dependent motors that transport membrane vesicles and various protein complexes along microtubules.

Microtubules Are Hollow Tubes Formed from Tubulin 14

Microtubules are formed from molecules of tubulin, each of which is a heterodimer consisting of two closely related and tightly linked globular polypeptides called α-tubulin and β-tubulin. Although tubulin is present in virtually all eucaryotic cells, the most abundant source for biochemical studies is the vertebrate brain. Extraction procedures yield 10 to 20% of the total soluble protein in brain as tubulin, reflecting the unusually high density of microtubules in the elongated processes of nerve cells.

Tubulin molecules themselves are diverse. In mammals there are at least six forms of α-tubulin and a similar number of forms of β-tubulin, each encoded by a different gene. The different forms of tubulin are very similar, and they will generally co-polymerize into mixed microtubules in the test tube, although they can have distinct locations in the cell and perform subtly different functions. The microtubules in six specialized touch-sensitive neurons in the nematode Caenorhabditis elegans, for example, contain a specific form of β-tubulin, and mutations in the gene for this protein result in the specific loss of touch-sensitivity with no apparent defect in other cell functions.

A microtubule can be regarded as a cylindrical structure in which the tubulin heterodimers are packed around a central core, which appears empty in electron micrographs. More accurately, perhaps, one can view the structure as being built from 13 linear protofilaments, each composed of alternating α- and β-tubulin subunits and bundled in parallel to form a cylinder ( Figure 16-21). Since the 13 protofilaments are aligned in parallel with the same polarity, the microtubule itself is a polar structure, and it is possible to distinguish a plus (fast-growing) and a minus (slow-growing) end.

Figure 16-21. Microtubules.

Figure 16-21

Microtubules. (A) Electron micrograph of a microtubule seen in cross-section, with its ring of 13 distinct subunits, each of which corresponds to a separate tubulin molecule (an α/β heterodimer). (B) Cryoelectron micrograph of a microtubule assembled (more...)

Microtubules Are Highly Labile Structures That Are Sensitive to Specific Antimitotic Drugs 15

Many of the microtubule arrays in cells are labile and depend on this lability for their function. One of the most striking examples is the mitotic spindle, which forms after the cytoplasmic microtubules disassemble at the onset of mitosis. The mitotic spindle is the target of a variety of specific antimitotic drugs that act by interfering with the exchange of tubulin subunits between the microtubules and the free tubulin pool. One of these is colchicine( Figure 16-22), an alkaloid extracted from the meadow saffron that has been used medicinally in the treatment of gout since ancient Egyptian times. Each molecule of colchicine binds tightly to one tubulin molecule and prevents its polymerization, but it cannot bind to tubulin once the tubulin has polymerized into a microtubule. The exposure of a dividing cell to colchicine, or to the closely related drug colcemid, causes the rapid disappearance of the mitotic spindle, indicating that a chemical equilibrium is maintained through continual exchange of subunits between the spindle microtubules and the pool of free tubulin. Because the temporary disruption of spindle microtubules preferentially kills many abnormally dividing cells, antimitotic drugs, such as vinblastine and vincristine (whose effects are similar to those of colcemid), are widely used in the treatment of cancer.

Figure 16-22. Chemical structures of colchicine and taxol.

Figure 16-22

Chemical structures of colchicine and taxol. A third drug, colcemid, is a close relative of colchicine in which the group shown in yellow is replaced by -CH3. Its binding to tubulin, unlike that of colchicine, is readily reversible.

The drug taxol ( Figure16-22), extracted from the bark of yew trees, has the opposite effect. It binds tightly to microtubules and stabilizes them, and when added to cells, it causes much of the free tubulin to assemble into microtubules. The stabilization of microtubules by taxol arrests dividing cells in mitosis, indicating that microtubules must be able not only to polymerize but also to depolymerize during mitosis. Taxol is also widely used as an anticancer drug.

Elongation of a Microtubule Is Rapid, Whereas the Nucleation of a New Microtubule Is Slow 16

Microtubule polymerization and depolymerization are complex and interesting processes with important biological roles. Most of what we know about the dynamic behavior of microtubules has come from studying the polymerization of purified tubulin molecules in vitro. Pure tubulin will polymerize into microtubules at 37°C in a test tube as long as Mg2+ and GTP are present. If the polymerization is followed either by light-scattering measurements or by microscopy, it shows an initial lag phase, after which microtubules form rapidly until a plateau level of polymerization is reached. The lag phase occurs because it is much easier to add subunits to an existing microtubule, a process called elongation, than to start a new microtubule de novo, a process called nucleation.

During the rapid polymerization phase, the high concentration of free tubulin causes microtubules to polymerize faster than they depolymerize (see below). When the plateau of polymerization is reached, however, not all of the tubulin will have polymerized because subunits are dissociating (depolymerizing) from the ends of microtubules as well as adding to them. The rate of polymerization drops with time because this rate is proportional to the concentration of free tubulin; the final concentration of free tubulin at the plateau, where the polymerization and depolymerization rates are exactly balanced, is called the critical concentration ( Figure 16-23).

Figure 16-23. Polymerization of pure tubulin.

Figure 16-23

Polymerization of pure tubulin. A mixture of tubulin, buffer, and GTP is warmed to 37°C at time zero. The amount of microtubule polymer, measured by light-scattering, follows a sigmoidal curve. During the lag phase individual tubulin molecules (more...)

We saw at the beginning of the chapter that the microtubules in a cell usually grow from a specific nucleating site (in most cases, the centrosome); because of a kinetic barrier to nucleation in solution, tubulin polymerization occurs only at this site. As in the test tube, not all the tubulin in the cell becomes polymerized. A typical fibroblast cell contains approximately 20 micromolar tubulin (2mg/ml), of which 50% is in microtubules and 50% is free.

The Two Ends of a Microtubule Are Different and Grow at Different Rates 17

The structural polarity of a microtubule, which reflects the regular orientation of its tubulin subunits, makes the two ends of the polymer different in ways that have a profound effect on its rate of growth. If purified tubulin molecules are allowed to polymerize for a short time at the ends of fragments of stable microtubules and the mixture is then examined in the electron microscope, one end can be seen to elongate at three times the rate of the other ( Figure16-24). The fast-growing end is thereby defined as the plus end and the other as the minus end.

Figure 16-24. Electron micrograph showing preferential polymerization of tubulin onto the plus ends of microtubules.

Figure 16-24

Electron micrograph showing preferential polymerization of tubulin onto the plus ends of microtubules. A stable bundle of microtubules obtained from the core of a cilium (discussed later) was incubated with tubulin subunits under polymerizing conditions. (more...)

It is possible to detect the polarity of microtubules in cross-section by adding free tubulin molecules to existing microtubules: under special conditions the tubulin monomers, instead of adding to the ends of the microtubules, add to the sides, forming curved protofilament sheets. In cross-section the sheets resemble hooks and, depending on the orientation of the microtubule, will appear to point either clockwise or counterclockwise ( Figure16-25). In this way it has been shown that the plus ends of the microtubules in a cell extend away from microtubule-nucleating sites such as the centrosome, the poles of a mitotic spindle, or the basal body of a cilium ( Figure 16-26).

Figure 16-25. Microtubule polarity as revealed by the hook-decoration method.

Figure 16-25

Microtubule polarity as revealed by the hook-decoration method. All the microtubules in this electron micrograph (seen in cross-section) have the same orientation. The hooks formed by the added tubulin curve clockwise, which indicates that the microtubules (more...)

Figure 16-26. The orientation of microtubules in cells.

Figure 16-26

The orientation of microtubules in cells. The minus ends of microtubules are generally embedded in a microtubule-organizing center, while the plus ends are often located near the plasma membrane.

Centrosomes Are the Primary Site of Nucleation of Microtubules in Animal Cells 18

The microtubules in the cytoplasm of an interphase cell in culture can be visualized by staining the cell with fluorescent anti-tubulin antibodies after the cells have been fixed. The microtubules are seen in greatest density around the nucleus and radiate out into the cell periphery in fine lacelike threads ( Figure16-27). The origin of the microtubules is seen most clearly if they are first depolymerized with colcemid and then allowed to repolymerize after the drug is washed out. The new microtubules grow out from the centrosome to form a small starlike structure called an aster and then elongate toward the cell periphery until the original microtubule distribution is reestablished ( Figure16-28). If the microtubules in cultured cells are decorated with tubulin hooks to determine their polarity, they are all seen to have their plus ends facing away from the centrosome, indicating that this organizing center has the capacity to nucleate microtubule polymerization with a specific polarity.

Figure 16-27. The interphase array of microtubules in a cultured fibroblast.

Figure 16-27

The interphase array of microtubules in a cultured fibroblast. The microtubules ( green) are stained with an antibody to tubulin; the cell nucleus ( blue) is stained with a fluorescent DNA-binding dye. (Courtesy of Nancy L. Kedersha.)

Figure 16-28. Microtubules growing out from the centrosome after the removal of colcemid.

Figure 16-28

Microtubules growing out from the centrosome after the removal of colcemid. Immunofluorescence micrographs showing the arrangement of microtubules in cultured cells as revealed by staining with anti-tubulin antibodies. A normal tissue-culture cell is (more...)

The centrosome is the major microtubule-organizing center in almost all animal cells. In interphase it is typically located to one side of the nucleus, close to the outer surface of the nuclear envelope. Embedded in the centrosome is a pair of cylindrical structures arranged at right angles to each other in an L-shaped configuration. These are centrioles, and we discuss their structure later. 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 (see Figure 18-5).

Surrounding each centriole pair, in both interphase and metaphase, is a region of the cytoplasm that stains darkly when viewed by electron microscopy and appears in the best micrographs to be made of a network of small fibers ( Figure 16-29A). This is the pericentriolar material, or centrosome matrix, and it is the part of the centrosome that nucleates microtubule polymerization. The protein composition of the centrosome matrix is only partly known, as is the mechanism by which it nucleates microtubules. However, it contains a number of centrosome-specific proteins, including a special minor form of tubulin, called γ-tubulin ( Figure 16-29B), which may interact with the normal α/β tubulin dimer to help nucleate microtubules.

Figure 16-29. The centrosome matrix.

Figure 16-29

The centrosome matrix. (A) Electron micrograph of a centrosome in a purified preparation. The matrix surrounds a barrel-shaped centriole, and it appears as a fibrous material that contains fine granules. (B) Light micrograph of a dividing human cell in (more...)

Not all microtubule-organizing centers contain centrioles. In mitotic cells of higher plants, for example, the microtubules terminate in poorly defined regions of electron density that are completely devoid of centrioles. Similarly, centrioles are not present in the meiotic spindle of mouse oocytes, although they appear later in the developing embryo. In fungi and diatoms the microtubule-organizing center is a plaque called the spindle pole body, which is embedded in the nuclear envelope. Despite these morphological differences ( Figure16-30), all of the organizing centers contain a matrix that nucleates microtubule polymerization, and they usually contain gamma-tubulin and other centrosome-specific proteins. Thus the molecular mechanism of microtubule nucleation is likely to be highly conserved.

Figure 16-30. A microtubule-organizing center in a fungal cell.

Figure 16-30

A microtubule-organizing center in a fungal cell. Electron micrograph of the spindle pole body in yeast. (Courtesy of John Kilmartin.)

Microtubules Depolymerize and Repolymerize Continually in Animal Cells 19

In a cell such as a cultured fibroblast the entire microtubule array is turning over rapidly. The half-life of an individual microtubule is about 10 minutes, while the average lifetime of a tubulin molecule, between its synthesis and proteolytic degradation, is more than 20 hours. Thus each tubulin molecule will participate in the formation and dismantling of many microtubules in its lifetime, a process that can be investigated by direct observation of living cells. One way is to inject tubulin that has been covalently linked to a fluorescent dye and then follow the behavior of microtubules that incorporate the tagged tubulin using fluorescence microscopy. Alternatively, in certain very flat cells one can visualize microtubules directly, without labeling them, using video-enhanced differential-interference-contrast microscopy (see Figure4-12). When microtubules in a cell are watched over time by either method, a remarkable phenomenon is observed. Individual microtubules grow toward the cell periphery at a constant rate for some period and then suddenly shrink rapidly back toward the centrosome. They may shrink partially and then recommence growing, or they may disappear completely, to be replaced by a different microtubule ( Figure16-31). These fluctuations in length occur over many micrometers and involve the polymerization and then depolymerization of tens of thousands of tubulin subunits. Transitions between prolonged periods of polymerization and depolymerization are also seen when pure microtubules are studied in a test tube ( Figure16-32). This behavior, called dynamic instability, plays a major role in positioning microtubules in the cell, as we discuss below.

Figure 16-31. Microtubule dynamics in a living cell.

Figure 16-31

Microtubule dynamics in a living cell. A fibroblast was injected with tubulin that had been covalently linked to rhodamine, so that approximately 1 tubulin subunit in 10 in the cell was labeled with a fluorescent dye. The fluorescence at an edge of the (more...)

Figure 16-32. The dynamic instability of microtubule growth.

Figure 16-32

The dynamic instability of microtubule growth. Fluctuations in length of a single microtubule in a solution of pure tubulin as seen by video-enhanced dark-field microscopy. Images of the same microtubule were recorded at intervals of 1 to 2 minutes and (more...)

GTP Hydrolysis Can Explain the Dynamic Instability of Individual Microtubules 20

The dynamic instability of microtubules requires an input of energy to shift the chemical balance between polymerization and depolymerization - energy that comes from the hydrolysis of GTP. GTP binds to the β-tubulin subunit of the heterodimeric tubulin molecule, and when a tubulin molecule adds to the end of a microtubule, this GTP molecule is hydrolyzed to GDP. (The α-tubulin subunit also carries GTP, but this cannot be exchanged for free GTP and is not hydrolyzed, so we can consider it a fixed part of the tubulin protein structure.)

The role of GTP hydrolysis in microtubule polymerization has been examined using analogues of GTP that cannot be hydrolyzed. Tubulin molecules containing such nonhydrolyzable GTP analogues form microtubules normally, indicating that, while the binding of this nucleotide is required for microtubule polymerization, its hydrolysis is not. These microtubules, however, are abnormally stable and do not depolymerize like normal microtubules when the tubulin concentration in the surrounding fluid is lowered or when they are treated with colchicine. Thus the normal role of GTP hydrolysis is apparently to allow microtubules to depolymerize by weakening the bonds between tubulin subunits in the microtubule.

Dynamic instability is thought to be a consequence of the delayed hydrolysis of GTP after tubulin assembly. When a microtubule grows rapidly, tubulin molecules add to a polymer end faster than the GTP they carry can be hydrolyzed. This results in the presence of a GTP capon the end of the microtubule, and because tubulin molecules carrying GTP bind to one another with higher affinity than tubulin molecules carrying GDP, the GTP cap will encourage a growing microtubule to continue growing. Conversely, once a microtubule has lost its GTP cap - for example, if the instantaneous rate of polymerization slows down - it will start to shrink and then tend to go on shrinking.

A model for the structural changes that accompany dynamic instability is shown schematically in Figure 16-33. Some general principles that apply to the polymerization of both actin filaments and microtubules are discussed in Panel 16-1, pages 824-825.

Figure 16-33. GTP hydrolysis after polymerization destabilizes microtubules.

Figure 16-33

GTP hydrolysis after polymerization destabilizes microtubules. Analysis of the growth and shrinkage of microtubules in vitro suggests the following model for dynamic instability. (A) Addition of tubulin heterodimers carrying GTP to the end of a protofilament (more...)

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Panel 16-1

The polymerization of actin and tubulin.

Cells can modify the dynamic instability of their microtubules for specific purposes. In each M phase of the cell cycle, for example, the rapidity with which microtubules form and break down is greatly increased, so that the chromosomes can readily capture growing microtubules and a mitotic spindle can rapidly assemble (discussed in Chapter 18). Conversely, when a cell differentiates and takes on a defined morphology, the dynamic instability of its microtubules is often suppressed by proteins that bind to the microtubules and stabilize them against depolymerization. The ability to stabilize microtubules in a particular configuration provides an important mechanism by which a cell can organize its cytoplasm.

The Dynamic Instability of Microtubules Provides an Organizing Principle for Cell Morphogenesis 21

Cytoplasmic microtubules in animal cells tend to radiate out in all directions from the centrosome, where their minus ends are anchored. Most animal cells are polarized, however, and the assembly and disassembly of tubulin molecules are spatially controlled so that microtubules extending toward specific regions of the cell predominate. It is not known for certain how this is achieved, but it seems likely that the mechanisms depend on the dynamic instability of microtubules.

We have seen that individual microtubules in vitro tend to exist in one of two states - steady growth or rapid, "catastrophic" disassembly - and that microtubules in a cell can also exist in these two states. The inherent instability of microtubules helps to explain how they can become organized in specific directions in a cell - toward the leading edge of a crawling cell, for example. The array of microtubules radiating from the centrosome is continually changing as new microtubules grow and replace others that have depolymerized. A microtubule that grows from a centrosome can be stabilized if its plus end is somehow stabilized, or capped, so as to prevent its depolymerization. If capped by a structure in a particular region of the cell, it will establish a relatively stable link between that structure and the centrosome. Microtubules originating in the centrosome can thus be selectively stabilized by events elsewhere in the cell. Cell polarity is thought to be determined in this way by unknown structures or factors localized in particular regions of the cell cortex that "capture" the plus ends of microtubules ( Figure 16-34).

Figure 16-34. The selective stabilization of microtubules can polarize a cell.

Figure 16-34

The selective stabilization of microtubules can polarize a cell. A newly formed microtubule will persist only if both of its ends are protected from depolymerizing. In cells the minus ends of microtubules are generally protected by the organizing centers (more...)

In many cells the initial stabilization of microtubules at their plus ends is consolidated to produce a more permanent polarization of the cell, as we now discuss.

Microtubules Undergo a Slow "Maturation" Revealed by Posttranslational Modifications of Their Tubulin 22

Tubulin subunits can be covalently modified after they polymerize. Two such modifications are especially interesting in that they provide a form of molecular clock, which can be used to tell how long it has been since a given microtubule polymerized. These modifications are the acetylation of α-tubulin on a particular lysine and the removal of the tyrosine residue from the carboxyl terminus of α-tubulin. Acetylation and detyrosination are both relatively slow enzymatic reactions that occur only on microtubules and not on free tubulin molecules; moreover, they are rapidly reversed as soon as a tubulin molecule depolymerizes. Thus the longer the time that has elapsed since a particular microtubule polymerized, the higher will be the fraction of its subunits that are acetylated and detyrosinated. Complete modification takes several hours, so that in fibroblasts, where microtubules turn over rapidly, relatively few of them are modified. In nerve axons, by contrast, the majority of microtubules are stable and most are modified.

Acetylation and detyrosination can be detected by specific antibodies, and they provide a useful indication of the stability of microtubules in cells in which it is difficult to study microtubule dynamics directly. The role of these modifications is unknown, but it is thought that they provide sites for the binding of specific microtubule-associated proteins that further stabilize mature microtubules.

Microtubule-associated Proteins (MAPs) Bind to Microtubules and Modify Their Properties 23

Whereas the posttranslational modification of tubulin marks certain microtubules as "mature" and may promote their stability, the most far-reaching and versatile modifications of microtubules are those conferred by the binding of other proteins. These microtubule-associated proteins, or MAPs, serve both to stabilize microtubules against disassembly and to mediate their interaction with other cell components. As one might expect from the diverse functions of microtubules, there are many kinds of MAPs; some are widely distributed in most cells, whereas others are found only in specific cell types.

Two major classes of MAPs can be isolated from brain in association with microtubules: HMW proteins (high-molecular-weight proteins), which have molecular weights of 200,000 to 300,000 or more and include MAP-1 and MAP-2; and tau proteins, which have molecular weights of 55,000 to 62,000. Proteins in both classes have two domains, only one of which binds to microtubules; the other is thought to help link the microtubule to other cell components ( Figure 16-35). Because the microtubule-binding domain binds to several unpolymerized tubulin molecules simultaneously, these MAPs speed up the nucleation step of tubulin polymerization in vitro. More important, they inhibit the dissociation of tubulin from the microtubule ends and thus stabilize the microtubules once they have formed. Staining with antibodies to MAP-2 and tau shows that both proteins bind along the entire length of cytoplasmic microtubules.

Figure 16-35. A microtubule-associated protein.

Figure 16-35

A microtubule-associated protein. (A) Electron micrograph showing the regularly spaced side arms formed on a microtubule by a large microtubule-associated protein (known as MAP-2) isolated from vertebrate brain. Portions of the protein project away from (more...)

Many other MAPs have been isolated. Some act as structural components and provide permanent links to other cell components, including other parts of the cytoskeleton. Others are microtubule motors, which use the energy of ATP hydrolysis to move along microtubules, as we discuss below.

MAPs Help Create Functionally Differentiated Cytoplasm 24

Many cell types specifically stabilize microtubules in specialized regions of cytoplasm. An especially well-studied example is provided by nerve cells, which extend two kinds of processes axons and dendrites. Axons, which are uniform in diameter and can be many centimeters long, are responsible for propagating electrical signals away from the cell body, whereas dendrites, which taper away from the cell body and rarely exceed 500 µm in length, are responsible for receiving electrical information from other neurons and relaying it to the cell body. Most nerve cells form several dendrites but only a single axon (see Figure 11-20).

Axons and dendrites are both packed with microtubules, although with different arrangements. In axons microtubules are very long and are all oriented with their plus ends away from the cell body. In dendrites the microtubules are shorter and their polarity is mixed: some have their plus ends pointing away from the cell body, while others have their plus ends pointing toward the cell body. When the distribution of MAPs in cultured neurons is studied with specific antibodies, certain forms of the tau protein are found to be present only in axons; MAP-2, on the other hand, is present in both dendrites and the cell body but completely excluded from axons ( Figure 16-36). Axons and dendrites are different in many other ways as well: mRNAs, ribosomes, and some kinds of ion channels, for example, are present in dendrites and the cell body but are excluded from axons, while certain cell-adhesion molecules and the Na+ channels involved in the generation of action potentials are selectively localized to axons. Thus both the cytoplasm and the plasma membrane of a nerve cell are divided into axonal and dendritic compartments. These compartments within a single cell differ from membrane-bounded compartments such as the endoplasmic reticulum or mitochondria, since they are not separated from each other by a membrane; instead, the difference seems to be one of structural organization and the types of proteins present.

Figure 16-36. An example of the cytoplasmic compartmentalization of nerve cells.

Figure 16-36

An example of the cytoplasmic compartmentalization of nerve cells. This micrograph shows the distribution of tau protein ( green) and MAP-2 ( orange) in a hippo-campal neuron in culture. Whereas tau is confined to the axon, MAP-2 is confined to the cell (more...)

The generation of axons and dendrites during the differentiation of nerve cells is discussed in Chapter 21. Although it is unclear how the cytoplasm and plasma membrane of a nerve cell become compartmentalized, MAPs may be essential for this process. When the production of tau protein is inhibited in cultured neurons by treatment with specific antisense oligonucleotides, the formation of axons is suppressed, whereas the formation of dendrites is unaffected. Conversely, when nonneuronal cells are genetically manipulated so that they express tau protein (which is normally expressed only in nerve cells), they form long axonlike processes, which contain bundles of microtubules arranged with their plus ends pointing away from the cell body, just as in nerve cells.

Because different components of the cell move along microtubules in different directions, one can postulate that an initial difference in microtubule polarity is created by a different distribution of MAPs, which will in turn lead to further differences between dendrites and axons. Secretory vesicles, for example, move toward the plus end of microtubules and therefore will be carried down the axon to the nerve terminals where they function; conversely, if ribosomes and mRNAs move toward the minus end of microtubules, they could be excluded from axons.

Kinesin and Dynein Direct Organelle Movement Along Microtubules 25

Important advances in cell biology have often followed the introduction of a new experimental technique, and it was the improved ability to see small faint objects by video-enhanced light microscopy that led to the discovery of the microtubule motors responsible for organelle transport. Once it became possible to visualize single microtubules in an unfixed specimen, investigators could follow the movement of organelles and other particles along these microtubules in vitro. Alternatively, they could observe and measure the gliding movement of individual microtubules over glass surfaces coated with cell extracts.

Such in vitro motility assays were used to identify and isolate two classes of microtubule-dependent motor proteins - the kinesins and the cytoplasmic dyneins. Cytoplasmic dyneins are involved in organelle transport and mitosis and are closely related to ciliary dynein,the motor protein in cilia and flagella (discussed later). Kinesins are more diverse than the dyneins, and different family members are involved in organelle transport, in mitosis, in meiosis, and in the transport of synaptic vesicles along axons. Both the cytoplasmic dyneins and the kinesins are composed of two heavy chains plus several light chains. Each heavy chain contains a conserved, globular, ATP-binding head and a tail composed of a string of rodlike domains. The two head domains are ATPase motors that bind to microtubules, while the tails generally bind to specific cell components and thereby specify the type of cargo that the protein transports ( Figure16-37).

Figure 16-37. Microtubule motor proteins.

Figure 16-37

Microtubule motor proteins. Kinesins and cytoplasmic dyneins are microtubule motor proteins that generally move in opposite directions along a microtubule (A). These proteins (drawn here to scale) are complexes composed of two identical heavy chains plus (more...)

The Rate and Direction of Movement Along a Microtubule Are Specified by the Head Domain of Motor Proteins 26

Most known motor proteins move in only one direction along microtubules - either toward the plus end or toward the minus end. This directionality can be analyzed in vitro by allowing polystyrene beads coated with the motor protein to move along microtubules that have been polymerized on centrosomes. Because the microtubules in such arrays have their plus ends outermost, the direction of movement can be readily determined with a light microscope. Whereas polystyrene beads coated with crude extracts of cytoplasm move in both directions, beads coated with kinesin isolated from axons move only outward toward the plus end of the microtubules. Beads coated with cytoplasmic dyneins, by contrast, move toward the minus ends of the microtubules, which are embedded in the centrosome.

Studies of intact nerve axons have confirmed the results obtained in in vitro experiments: organelle movement away from the cell body is driven mainly by kinesin, whereas organelle movement back from the nerve terminal toward the cell body is driven by cytoplasmic dynein ( Figure16-38). Since all proteins are made in the nerve cell body, cytoplasmic dynein must be carried first in a nonfunctional state to the nerve terminal before it can begin to work to transport organelles back to the cell body.

Figure 16-38. Vesicle transport in two directions.

Figure 16-38

Vesicle transport in two directions. Kinesin and cytoplasmic dynein carry their cargo in opposite directions along microtubules, as illustrated in a fibroblast (A) and in the axon of a neuron (B).

Surprisingly, not all kinesins move organelles toward the plus end of microtubules. A Drosophila kinesin called Ncd, for example, which is required for normal meiosis, differs from axonal kinesin in both the direction and the rate at which it moves along microtubules: whereas axonal kinesin walks toward the plus end at approximately 2 µm/second, the Ncd protein walks toward the minus end at about 0.1 µm/second.

The mechanism by which these motor proteins convert the energy of ATP hydrolysis into vectorial movement is not known. Finding out how two closely related head domains can move in opposite directions along a microtubule will require detailed structural studies and is likely to illuminate the energy transduction process itself.


Microtubules are stiff polymers of tubulin molecules. They assemble by addition of GTP-containing tubulin molecules to the free end of the microtubule, with one end (the plus end) growing faster than the other. Hydrolysis of the bound GTP takes place after assembly and weakens the bonds that hold the microtubule together. Slowly growing microtubules are especially unstable and liable to catastrophic disassembly, but they can be stabilized in cells by association with other structures that cap their two ends. Microtubule-organizing centers such as centrosomes protect the minus ends of microtubules and continually nucleate the formation of new microtubules, which grow out in random directions. Any microtubule that happens to encounter a structure that stabilizes its free plus end will be selectively retained, while other microtubules will depolymerize. It is thought that this selective process largely determines the position of the microtubule arrays in a cell.

The tubulin subunits in microtubules that have been selectively stabilized are modified by acetylation and detyrosination. These alterations are thought to label the microtubule as "mature" and provide sites for the binding of specific microtubule-associated proteins (MAPs), which further stabilize the microtubule against disassembly. Microtubule motor proteins constitute an important class of MAPs that use the energy of ATP hydrolysis to move unidirectionally along a microtubule, carrying specific cargo. In general, dyneins move cargo toward the minus ends of microtubules, while most kinesins move cargo toward the plus ends. Such motor proteins are largely responsible for the spatial organization and directed movements of organelles in the cytoplasm.

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Copyright © 1994, Bruce Alberts, Dennis Bray, Julian Lewis, Martin Raff, Keith Roberts, and James D Watson.
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