Microtubules and microtubule

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Microtubules and microtubule-associated proteins
Eckhard Mandelkow and Eva-Maria Mandelkow
M a x Planck Unit for Structural Molecular Biology, Hamburg, Germany
Microtubule research is becoming increasingly diverse, reflecting the many
isoforms and modifications of tubulin and the many proteins with which
microtubules interact. Recent advances are particularly visible in four areas:
microtubule motor proteins (their structures, stepping modes, and forces);
microtubule nucleation (the roles of centrosomes and y-tubulin); tubulin
folding (mediated by cytoplasmic chaperones); and the expanding list of
microtubule-associated proteins, knowledge of their phosphorylation states,
and information on their effects on microtubule dynamics.
Current Opinion in Cell Biology 1995, 7:72-81
Introduction
Microtubules represent one of the fiber systems of the
eukaryotic cytoskeleton. They are essential for a wide
variety of cellular functions, notably cell motility, transport, cell shape and polarity, and mitosis. Microtubules
consist of a core cylinder built from heterodimers of etand ~-tubulin monomers. Three main classes of proteins
interact with tubulin. The 'inicrotubule-associated proteins', or MAPs, are sometimes called more specifically
'structural' MAPs because they bind to, stabilize and promote the assembly ofmicrotubules, and because they can
be copurified with tubulin through several cycles of
microtubule assembly and disassembly. Representatives
are MAPla and lb, MAP2a, 2b, and 2c, MAP4, tau
protein, 205 kDa MAP, and isoforms of these proteins
that are often generated by alternative splicing.
Another broad class comprises the motor proteins, so
called because they generate movement along microtubules using the chemical energy of ATP hydrolysis.
Representatives are kinesin (a motor moving towards the
distal, or plus, end of microtubules), dynein (a rearward
motor, also involved in the bending of cilia and flagella),
and their many relatives. The motors can bind tightly to
microtubules, for example in the absence ofnucleotides,
but they do not copurify through cycles of assembly and
disassembly.
A third and more heterogeneous class includes proteins that are not normally called MAPs but are often
found associated with microtubules and inay even copurify with them. Examples are glycolytic enzymes (e.g.
GAPDH and aldolase), kinases (e.g. protein kinase A,
GSK-3 and c-mos), proteins involved in biosynthesis
(elongation factor EF-lCt and even entire ribosomes),
proteins linking up to membrane receptors (dynamin and
G proteins), and ribonucleoproteins carrying mKNA.
These interactions are not always well defined, but they
point to the cytoskeleton as a (transient) anchor for many
cytoplasmic proteins.
The fields of microtubules, MAPs, and motors have expanded so nmch that they have become the subject of
separate reviews. For example, microtubule structure,
assembly and regulation have been reviewed recently
[1-4], as have MAPs [5,6] and motors [7-9]. General
surveys of microtubules and related proteins are given in
[10,11].
This review covers selected articles concerning microtubules, MAPs and motor proteins published in the last
eighteen months.
Structural aspects of microtubules revealed by
motors
The information on microtubule structure we have today
is derived mostly from electron microscopy combined
with image reconstruction, X-ray scattering, and video
microscopy; information on tubulin structure is indirect and derived mostly from biochemical experiments
(binding sites ofnucleotides or drugs, cross-linking studies, and so on). Novel insights on the microtubule lattice have recently come from studies of the interaction
between microtubules and kinesin. The head domain
of kinesin binds to [3-tubulin with a stoichiometry of
one kinesin head per tubulin heterodimer, generating an
axial periodicity of 8 nm (the height of the dimer) and
a 'B'-lattice in which adjacent protofilaments are staggered by about 0.9 nm [12°-14"]. The dimer polarity is
such that ~-tubulin points to the plus (fast-growing) end
of the microtubule and [3-tubulin points towards the minus (slow-growing) end (see Fig. 1). This implies that the
plus end has a crown ofet-tubulin subunits and the minus
end terminates with [3-tubulin [14°]. This polarity enables the ~-subunits to interact with T-tubulin, which is
a key component ofmicrotubule-organizing centers (reviewed in [3]). As the exchangeable GTP-binding site is
Abbreviations
GAPDH--glyceraldehyde-3-phosphate dehydrodgenase;GSK-3--glycogen synthasekinase-3; MAP~microtubule-associated protein.
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© Current Biology Ltd ISSN 0955-0674
Microtubules and microtubule-associated proteins Mandelkow and Mandelkow
on ~-tubulin and the microtubule-bound GTP is at the
plus end [15°], another consequence of this arrangement
of the microtubule is that the 'GTP-cap' sits on the plus
end in an inverted fashion so that the last ~-tubulin-GTP
is buried by a terminal layer of et-tubuhn (see Fig. 1).
region of ~-tubulin [39,40]. The stoichiometry of taxol
binding is one molecule per tubulin dimer; dimers stabilized by taxol are capable of assembling with bound
GDP, without the usual need for GTP hydrolysis [41].
Kinesin moves along the microtubule in 8 nm intervals
and produces forces in the pN range [16°-20"]. The direction of movement is parallel to the tracks of protofilaments [21], and the step size matches the separation
of the ~-tubulin 'stepping stones'. As kinesin works as
a dimer in vivo and shows alternating head catalysis (using each head of the dimer alternately to hydrolyze ATP)
[22°], one could integrate the structural and kinetic data
by assuming that a kinesin dimer would 'walk' or 'hop'
along one or two adjacent protofilaments. (Note that if
the center o f the kinesin molecule advances by 8 nm, this
may imply 16 nm steps for each head.) It also seems that
kinesin prefers rigid stepping stones over wobbly ones:
microtubules are fairly stiff structures, with persistence
lengths (a measure of their straightness) in the mm range
[23-25], but it is possible to make them even stiffer using
slowly hydrolyzable GTP analogues such as GMP-PCP,
in which case the movement of kinesin becomes 30%
faster [26].
GTP binding to tubulin
Microtubules contain a non-exchangeable GTP on
ct-tubulin and an exchangeable one on ~-tubulin (nucleotides on the latter site can be exchanged with nucleotides from the solution). Extending earlier work,
two recent studies showed that the exchangeable G T P
site is in the amino-terminal domain o f [~-tubulin; an
additional ATP-binding site (not involved in microtubule assembly) was found on et-tubulin [42,43]. That
microtubule stability is closely linked to GTP binding
and hydrolysis has now been demonstrated directly by
Davis et al. [44"], who made several mutant tubulins in
which altered GTPase activity is paralleled by altered
dynamic instability. Similarly, Reijo et al. [45 °] made
systematic mutations of clusters o f charged amino acids
for alanine in yeast [~-tubulin and deduced that three regions are critical for ~-tubulin function: near the carboxyl terminus, around residue 330, and around residue
150 (near the glycine cluster that is probably involved in
GTP binding).
The microtubule-motor interaction is surprisingly versatile: there are variants ofkinesin such as Drosophila ncd
or Saccharomyces cerevisiae Kar3 that can walk in the opposite direction (towards the minus end), and this property is inherent to the head domains [27°,28,29]. The
motor can twist its neck in order to attach to microtubules in the proper direction [30], and may even cause
rotation o f microtubules instead o f translation, as is suggested by the presence ofkinesin-like proteins on the C2
central microtubule in flagella in Chlamydomonas [31].
Finally, CENP-E, a kinesin-like protein that migrates
from kinetochores to the midzone of mitotic spindles,
cross-links the antiparallel microtubules and presumably
controls their rigidity or ghding during anaphase [32°].
Tubulin structure and interactions
Drugs that target tubulin
Tubulin has many applications as a target of drugs
(e.g. anti-cancer drugs) and therefore is a workhorse for
certain drug screening tests. The different modes of action and binding sites on tubulin of various drugs continue to be explored (see reviews [33,34]). Colchicine
and vinblastine are examples of drugs that destabilize
microtubules and thus disrupt mitosis; however, their effect can be felt even at drug concentrations too low to
cause net microtubule disassembly. This can be attributed
to a pronounced effect of the drugs on microtubule dynamic instability parameters [35-37], which are closely
linked to G T P binding and hydrolysis (see below). The
binding site for colchicine has been mapped near the
amino terminus of ~-tubulin [38]. By contrast, taxol
stabihzes microtubules, thereby also disrupting mitosis.
It was shown using photoactivatable taxol derivatives
that taxol binds predominantly to the amino-terminal
Modifications of tubulin
An enigmatic feature of tubulin is its heterogeneity. Not
only are et- and [~-tubulin each encoded by up to six
different genes, but the protein can also be modified
in several ways: phosphorylation, acetylation, detyrosination, and glutamylation. The complexity of neuronal
tubulins is particularly great and appears to increase with
differentiation. In other cells, such as avian erythrocytes,
the degree o f heterogeneity is conspicuously low [46].
One function of neuronal tubulin modification may be
the selective stabihzation of certain microtubules [47].
One of the modifying enzymes, the tubulin tyrosine
ligase, has now been sequenced and characterized in
detail [48]. This enzyme restores a carboxy-terminal
tyrosine to et-tubulin after it has been cleaved by a
tubulin carboxypeptidase. However, et-tubulin can also
lose two carboxy-terminal residues (glutamic acid and
tyrosine). This A2-tubulin can no longer be retyrosinated by the ligase and is enriched in very stable neuronal
microtubules [49]. Another sign of neuronal differentiation is the increasing length ofpoly-glutamic acid chains
attached to glutamic acid residues near the carboxyl terminus of 0t- and ~-tubulin; this makes the already very
acidic carboxyl termini even more negatively charged
[50]. The functions of these post-translational modifications have been ditlqcult to ascertain, but Gurland
and Gundersen [51] found that long-lived stable microtubules (known as 'Glu-microtubules' because they
have lost the carboxy-terminal tyrosine from ct-tubulin)
selectively break down when phosphatases are inhibited
by okadaic acid. One interpretation of this observation
is that stabilization factors (caps) that are normally associated with stable microtubules can be lost as a result of
unchecked phosphorylation.
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Cytoskeleton
Regulation of tubulin synthesis
The level of tubulin synthesis is regulated via the soluble
tubulin pool, whose size is in turn coupled to the degree
of microtubule assembly. In the case of ~-tubulin this
autoregulation involves the four amino-terminal residues
(MR_El, in the single letter code for amino acids) [52].
In an extension of their earlier work, Cleveland and coworkers [52] showed that 0t-tubulin follows a different
pathway (even though both subunits are downregulated
by degrading their mRNAs).
Prokaryote homologs of tubulin?
Tubulin is thought to be restricted to eukaryotic cells,
but there is a curious homology of the conserved
glycine-rich cluster (GGGTGSG) to the bacterial FtsZ
protein from Escherichia coll. Like tubulin, this protein
is also involved in cell division and binds GTP, and
Bramhill and Thompson [53] have now shown that
it even polymerizes in a GTP-dependent fashion, suggesting that the similarity is more than accidental, even
though the overall homology is very low.
Centrosomes and chaperones
(D 1995 Current Opinion in Cell Biology
cz-tubulin
O [3-tubulin-GTP
0 J3-tubulin-GDP
Fig. 1. Several features of the microtubule structure deduced from
labeling microtubules with the kinesin motor domain (for details,
see [14"]). The plus (fast-growing, distal) end is up, the minus
end (slow-growing, proximal) is down. The minus end is normally
buried in the microtubule-organizing structure containing y-tubulin (not shown here). An ct[B-tubulin heterodimer is shown near the
top (height 2 x 4 = 8 nm). The plus end of a microtubule terminates
with a crown of (~-tubulin, the minus end with a crown of ifl-tubulin. [3-Tubulin carrying GTP occurs in solution and at the plus end
[15"], but not in the interior of the microtubule. Note that the [3tubulin-GTP cap is covered by the terminal layer of ct-tubulin. The
protofilaments are shifted axially by 0.9 nm, corresponding to the
'B4attice' which results in a discontinuity in the microtubule wall
(moved to the back and thus not shown in this diagram). In the fully
saturated microtubule lattice every tubulin heterodimer binds one
kinesin head, thus emphasizing the dimer lattice of microtubules.
Native kinesin is a dimer of two heavy chains, linked via a flexible
neck and a coiled-coil stalk (simplified here as a V-shape), and two
light chains. The diagram shows three hypothetical modes of attaching the kinesin dimer to the microtubule: (a) on two adjacent
protofilaments side by side; (b) on one protofilament (kinesin heads
displaced axially by one tubulin dimer, or 8 nm); (c) on two adjacent protofilaments (heads staggered by one tubulin dimer). Each
of these modes would be compatible with the observed structure
of the microtubule decorated with kinesin.
Components of the centrosome
For microtubules to operate properly they must be nucleated at the right time and in the right place. This
normally happens at the centrosome, which consist of
a pair of centrioles surrounded by periocentriolar material; the latter is where microtubule nucleation actuary occurs. The stages and components of centrosome
assembly are beginning to be understood. One of the
components is 7-tubulin, which exists in the cytoplasm
as part of a multiprotein complex ('gammasome') with
several other proteins. 7-Tubulin is recruited to the centrosome for nucleation of microtubules at their minus
ends; nucleation is blocked when 7-tubulin is depleted or
inactivated. Other components of the centrosome precursor include ct- and ~-tubulin, actin, and the molecular chaperone hsp70 [54"-57"]. Additional proteins are
required to assemble the pericentriolar material, such
as pericentrin, an elongated 220kDa protein with a
coiled-coil stalk and two globular ends, identified using antisera from patients with scleroderma [58"]. This
general domain structure is shared with other proteins
found in microtubule-organizing structures, such as the
SP-H antigen/NuMa [59], and variants of tektin, an
(~-helical protein first found in flagellar axonemes and
recently in centrosomes [60]. The microtubule-nucleating function of 7-tubulin in centrosomes appears to
be highly conserved as human 7-tubulin functions even
when introduced into yeast [61].
Chaperones
Before microtubules can be nucleated, the tubulin must
first be folded; this is achieved by another oligomeric
complex, the cytoplasmic chaperone complex TCP-1,
Microtubules and microtubule-associated proteins Mandelkow and Mandelkow
also known as TriC. This 800 kDa structure contains several homologs of the heat shock protein Hsp60 (termed
CCT0t, -~, and so on) arranged in the two-layered
ring structure also found in bacterial and mitochondrial chaperones; this complex assists in the folding o f
relatively few proteins, among them the three tubulins
and actin [62"-64"]. TCP-1 subunits are the cytosolic
equivalents of the bacterial chaperone GroEL and mitochondrial Hsp60. The folding takes place in several steps
such that the initial association occurs with ADP bound
to the complex, followed by a later release involving
ATP hydrolysis [65"]. Yeast have equivalent chaperones
(BIN2, BIN3 and so on); even though they are related,
they appear to serve somewhat different functions and
do not replace one another [66"].
Location of assembly and transport
A related and more specialized problem is the nucleation
of microtubules in nerve cells, whose long processes are
filled with microtubules that serve as tracks for intracellular transport. Are the microtubules assembled in the
axon or in the cell body? The latter appears to be the
case: nucleation sites containing ~,-tubulin are found only
in the cell body, and microtubules are then transported
into the axons or dendrites (see below). Superimposed
on this transport process is a length redistribution which
allows microtubules to change their length, presumably
by dynamic instability, in the axon. The plus ends of the
transported microtubules are oriented distally, and this
end is where incorporation o f new subunits can occur
[67].
Microtubule-associated proteins
The majority of studies have traditionally been done
with brain MAPs, partly because brain tissue was the
most abundant source of microtubules, and partly because the MAPs provide selective markers for brain
development and cell type. The best known MAPs are
the heat-stable proteins MAP2 and tau; they are localized (mainly) to the dendritic and axonal compartment,
respectively, and they share homologous repeats in the
carboxy-terminal region that contribute to microtubule
binding (reviewed by Schoenfeld and Obar [51).
Tau
Tau is thought to stabilize microtubules in axons and
thus provide the basis for axonal transport. Consistent
with this idea, it was found that when tau is transfected
into culture cells microtubules become more stable and
the degree of assembly increases [68,69]. This effect can
be observed even more dramatically in insect Sf0 ceils
transfected with tau using the baculovirus system: these
normally round cells develop 'neurite-like' processes. In
this system, tau also protects microtubules against druginduced disassembly [70",71"]. The simple picture does
not always hold, however, as transfection oftau into Chinese hamster ovary cells did not result in an increase in
microtubule stability or mass [72"]. Even more surprising
is the fact that tau-null transgenic mice are viable; in this
case, the stabilization ofmicrotubules may be achieved by
other MAPs [73"].
Functions of tau
Tau has several distinct effects on microtubules in vitro:
it binds to them, promotes their nucleation and elongation, protects against disassembly, and induces them
to form parallel arrays (bundles). These functions can
be traced back to different domains of tau. For example, efficient assembly of microtubules requires not only
the internal repeat domain, but also regions on either
side, acting as 'jaws'; the flanking regions in turn tend
to promote the formation of bundles [74-76]. Bundles
of microtubules may be important not only in neurite outgrowth, but also in other functions such as in
the erythrocyte marginal band, where tau is one of the
MAPs responsible for cohesion [77]. Although the contribution o f the repeat domain to microtubule binding is
generally weak, there are certain 'hotspots' (e.g. the region between repeats 1 and 2) which could regulate the
affinity, especially during the developmentally regulated
alternative splicing o f tau [78].
Distribution
Another characteristic feature o f tau is its heterogeneous
distribution, both within cells and between different cell
populations. Whereas six tau isoforms are found in central nervous tissue (in humans), 'big' tau (distinguished
mainly by the presence o f an additional exon, 4a) occurs in peripheral nerves. Within neurons, tau is mainly
axonal (in contrast to the somatodendritic MAP2), but
there is also a nuclear variant of tau. This intracellular
sorting may take place at the level of the transcripts,
which come in sizes between 2.5 and 8 kilobases and may
contain targeting information. Microtubules may play a
role in distributing tau, because m R N A is transported
along them [79-82].
AIzheimer's disease
Apart from its cell biological role, tau has generated
considerable interest because of its relationship with
Alzheimer's disease. Tau is the main component of the
paired helical filaments that make up the neurofibrillary tangles in the brains of patients with Alzheimer's
disease. In these tangles, tau is not only abnormally
aggregated, but also modified in several other ways fieviewed in [83]). The most conspicuous modification is
excess phosphorylation, frequently at Set-Pro or ThrPro motifs (indicating the influence of proline-directed
kinases). There are other sites (such as Ser262) that
strongly influence the affinity of tau for microtubules
[84"] and are phosphorylated in the tau protein found in
Alzheimer's disease [85]; phosphorylation might explain
why this tau does not bind to microtubules and ceases
to stabilize them [86]. This effect contrasts with that resulting from 'fetal' phosphorylation of tau, which shares
some of the phosphorylation sites of Alzheimer's disease,
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Cytoskeleton
but which still binds well to microtubules [87]. Several
phosphorylation sites in fetal and adult rat tau have now
been determined directly by mass spectrometry and sequencing [88]. From a practical point of view, most studies of phosphorylation rely on phosphorylation-dependent antibodies, which can also be used with cell models
(epitopes are summarized in [83]). All of the sites found
so far can be dephosphorylated by phosphatases PP-2A
and PP-2B (calcineurin), both of which are abundant in
brain [89]. Other modifications include ubiquitination at
several lysines [90] and glycation [91].
Structure
How the normal and pathological properties o f tau are
related to its structure remains enigmatic. In solution, tau
lacks secondary structure or even a defined shape and is
best described as a random coil; the lack of secondary
structure is observed even in paired helical filaments,
suggesting that these may not assemble by way of interacting ~-strands, in contrast to other fibers found in
pathological states [92]. Nevertheless, tau tends to associate into antiparallel dimers, and in this form it is prone
to assembly into paired helical filaments [93].
MAP2, MAPla, MAPlb and MAP4
MAP2
MAP2 is a relative o f tau in that it has a homologous repeat domain near the carboxyl terminus but has a much
larger extension towards the amino terminus. The middle part can be spliced out to give MAP2c, a 'juvenile'
isoform [94]. Whereas tau can have either three or four
repeats, MAP2 used to be found with three repeats only;
however, this gap has now been closed by the discovery
o f a four-repeat MAP2c. Unlike the neuronal MAP2,
this isoform occurs in gila cells [94]. The question of
how MAPs bind to microtubules, and how they interfere
with motor proteins, continues to-be a matter of study.
Wallis et al. [95] showed that MAP2 binds in a positively cooperative manner, leading to the clustering of
MAP2 on the microtubule surface (which makes it distinct from tau [76]). Echoing an earlier debate, Hagiwara
et al. [96] reported that MAP2 and motor proteins compete for the same binding sites (in the carboxy-terminal
domain o f tubulin), whereas Marya et al. [97] found distinct sites and no competition. A compromise solution
was offered by Lopez and Sheetz [98], who found distinct sites but steric hindrance, as might be expected with
molecules as large as MAP2. Whatever the explanation
o f the results, in practice there seems to be no doubt that
motors can work their way along microtubules quite efficiently, even when MAPs are present.
M A P l a and M A P l b
MAPla and M A P l b are two other high molecular
weight neuronal MAPs; they do not belong to the
same class as MAP2 and tau because they have different microtubule interaction motifs. MAPla and M A P l b
share partial sequence homology, and both are syn-
thesized as a polyprotein coupled to a light chain
(LC; M A P l b - L C 1 , 34 kDa, and MAPla-LC2, 28 kDa,
respectively [99,100]). As M A P l b binds to microtubules
through a series of basic repeats o f the type KKE,
the same mode of binding had also been assumed for
MAPla. This is not the case, however, as Cravchik et al.
[101] have discovered that MAPla has a novel binding
motif which is unique, as it is acidic rather than basic.
They proposed that MAPla has a bipolar helical structure, with positive and negative charges separated on the
two sides. The third light chain, LC3 (18kDa), is encoded separately and has now been cloned [102]. It is
a basic protein, abundant in neurons, that binds to both
M A P l a and M A P l b and could modulate their interaction with microtubules.
MAP4
In contrast to the neuronal proteins discussed above,
MAP4 has a ubiquitous distribution. It resembles MAP2
and tau in having a homologous repeat domain that
interacts with microtubules. Contrary to the view that
MAPs are freely diffusible in cells, Chapin and Bulinski
[103] have observed a surprising extent o f microheterogeneity of MAP4-microtubule interactions in cell cultures, suggesting the selective stabilization of subsets of
microtubules within cells. A related MAP4-1ike protein
has been purified from Xenopus eggs [104]. This protein becomes phosphorylated during mitosis and thus is
a candidate for MAP4-dependent regulation of microtubule dynamics.
Plant MAPs
Compared to the proteins mentioned so far, the study of
plant microtubules has been lagging behind, partly because o f the difficulty in preparing sufficient quantities
o f protein. Recent work shows that plant microtubules
are quite comparable with those of animals; for example,
plant tubulin can be assembled reversibly using taxol, hybrid systems o f maize tubulin assembled with neuronal
MAP2 show tight binding (dissociation constant in the
micromolar range), and some plant MAPs appear to be
related to tau [105-107].
Other proteins associated with microtubules
As mentioned in the Introduction, not all associated proteins can be classified as structural or motor MAPs;
conversely, as microtubules are 'sticky', not all associations may be meaningful. As phosphorylation regulates
many intracellular interactions, it is intriguing that microtubules associate with a number ofkinases (such as the
mitogen-activated protein kinase cdk5, and GSK-3; see
[83]) that can cause the 'pathological' phosphorylation of
tau protein. The kinases often bind to microtubules via
MAPs. An interesting case is that o f protein kinase A associated with MAP2: phosphorylation of the regulatory
subunit of protein kinase A by the cell cycle kinase cdc2
releases the catalytic subunit and presumably allows it to
phosphorylate other targets [108]. Dynamin is a GTPase
involved in endocytosis; it has two microtubule-bind-
Microtubulesand microtubule-associatedproteinsMandelkow and Mandelkow
ing sites near the amino terminus and a proline-rich
domain (resembling Src homology region 3 domains)
near the carboxyl terminus whose affinity is regulated
by phosphorylation [109].
Several proteins involved in signal transduction bind
tightly to tubulin or microtubules. Recent examples include neurofibromin (a neuronal GTPase-activating protein) [110], the ~-subunits of several G proteins (which
can take up GTP from tubulin) [111], and gephyrin (a
protein linked to the glycine receptor) [112]. Associations with microtubules are not restricted to proteins;
the binding of mRNAs was mentioned above [80], and
recently Surridge and Burns [113] reported that the middie part of MAP2 has a high affinity for phospholipids,
a property not found in tau or MAP2c.
O f special interest in this context are proteins that are
capable ofmicrotubule severing, which could contribute
to their rapid reorganization, for example during mitosis. Examples include katanin, a heterodimeric protein
(60 and 81kDa) that severs microtubules in an ATPdependent fashion [114"], and elongation factor EF-I~,
a GTP-binding protein involved in protein synthesis
[115°]. On the other hand, a protein like EF-10t has
also been found to be able to cause the formation of
bundles of microtubules [116 °] and has been found in
the pro-centrosomal protein complex (see above; [54°]).
It will be interesting to see how these seemingly contradictory activities are related to one another. Finally,
flagellar microtubules can be excised near their base,
and it turns out that this is achieved by contractile fibers
consisting of centrin, a relative of the calcium-binding
protein calmodulin ([117"]; see also Salisbury, this issue,
pp 39-45).
Microtubule dynamics
One of the most fascinating aspects of microtubules is
their rapid turnover and potential for reorganization,
unexpected for a cyto-'skeletal' polymer. The half-lives
of most microtubules are in the range of minutes, and
the predominant mechanism of redistribution appears to
be dynamic instability (reviewed by Cassimeris, [118]),
perhaps in combination with the severing factors just
mentioned [114",115°]. Dynamic instability can also
be observed in plant cells [119]. On the other hand,
many cellular microtubules are intrinsically stable [120],
suggesting that the observed dynamics are tightly regulated by factors controlling catastrophes and rescues,
including kinases such as the cell cycle dependent kinase cdc2. Independent of these mechanisms is a steady
flux (or treadmill) of subunits, such that labeled subunits migrate slowly from the plus to the minus end
of the microtubule [121]. In spite of this activity, and
contrary to intuition, the assembled microtubule mass
does not change nmch during mitosis, indicating that
reorganization does not require net disassembly [122].
A reconstruction of an entire spindle microtubule assembly shows that the nearest-neighbor interactions are
mainly between antiparallel microtubules, supporting the
potential for bidirectional gliding during anaphase [123].
Plant ceils have the capacity for reorienting their entire
microtubule network in response to extracellular factors
and gravity [124]. Some of these features can be reproduced in vitro, where microtubules can be induced to
display elaborate waves of assembly and orientation in
time and space [125"].
Conclusions
In contrast to the actin-based cytoskeleton, where the
structures of the main players (actin, myosin and their
complex) have been solved by X-ray crystallography
[126,127], the microtubule system does not yet offer
such insights. A direct structural analysis may be possible by electron crystallography, because zinc induces
the formation of tubulin sheets that diffract to nearatomic resolution [128°]. For the time being, we have
to learn by analogy. For example, the structure of the
actin-binding domains of gelsolin [129] illustrates that
repetitive sequences need not reflect repetitive binding; the interactions may in fact be based on the variable
parts, whereas the repeats could simply provide a framework for protein folding (this argument might apply to
models of rnicrotubule-MAP interactions). Tubulin is a
GTP-binding protein that shares weak homology with
signal-transducing G proteins. The structure of the ctsubunit of transducin has become available [130]; part
of it folds in a manner similar to the small G protein
1Las/p21 (reviewed in [131]). Finally, if one is interested
in an example of an assembly of 0r- and [~-subunits, with
nucleotides bound in a non-exchangeable and exchangeable fashion, then the recent solution of the F1-ATPase
structure may be worth studying [132]. Even though
these proteins have different functions from tubulin, they
provide intriguing food for thought.
Acknowledgements
We thank Alexander Marx for help in compiling the references,
and Stefan Sack for the preparation of Fig. 1.
References and recommended reading
Papers of particular interest, published within the annual period of
review, have been highlighted as:
*
of special interest
**
of outstanding interest
1.
Guenette S, Solomon F: Microtubule assembly: pathways, dynamics, and regulators. Curr Opin Struct Bio1199 3, 3:198-20].
2.
Mandelkow E, Mandelkow E-M: Microtubule structure. Curr
Opin Struct Biol 1994, 4:171-179.
3.
Joshi HC: Microtubule organizing centers and y-tubulin. Curt
Opin Cell Biol 1994, 6:55-62.
4.
Hirokawa N: Microtubule organization and dynamics dependent on microtubule-associated proteins. Curt Opin Cell Biol
1994, 6:74-81.
5.
Schoenfeld TA, Obar RA: Diverse distribution and function of
fibrous microtubule-associated proteins in the nervous system.
Int Rev Cytol 1994, 151:67-137.
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Cytoskeleton
6.
Lee G: Non-motor microtubule-associated proteins. Curt Opin
Cell Biol 1993, 5:88-94.
A kinetic analysis of the interactions between microtubules, kinesin, and
ATP hydrolysis.
7.
Goldstein LSB: With apologies to Scheherazade: tails of 1001
kinesin motors. Annu Rev Genet 1993, 27:319-351.
23.
8.
Walker R, Sbeetz MP: Cytoplasmic microtubule-associated motors. Annu Rev Biochem 1993, 62:429-451.
Dye RB, Fink SP, Williams RC: Taxol-induced flexibility of microtubules and its reversal by MAP2 and tau. J Biol Chem
1993, 268:6847-6850.
24.
Gittes F, Mickey B, Nettleton J, Howard J: Flexural rigidity
of microtubules and actin filaments measured from thermal
fluctuations in shape. J Cell Biol 1993, 120:923-934.
25.
Venier P, Maggs AC, Carlier MF, Pantaloni D: Analysis of microtubule rigidity using hydrodynamic flow and thermal fluctuations. J Biol Chem 1994, 269:13353-13360.
26.
Vale RD, Coppin CM, Malik F, Kull FJ, Milligan RA: Tubulin
GTP hydrolysis influences the structure, mechanical properties, and kinesin driven transport of microtubules. J Biol Chem
1994, 269:23769-23775.
9.
SkoufiasDA, Scholey JM: Cytoplasmic microtubule-based motor proteins. Curr Opin Cell Biol 1993, 5:95-] 04.
10.
Kreis 1-, Vale R (Eds): Guidebook to the cytoskeletal and motor
proteins. Oxford: Oxford University Press; 1993.
11.
Hyams JS, Lloyd CW (Eds): Microtubules. New York: Wiley-Liss;
1993.
12.
•
Song Y-H, Mandelkow E: Recombinant kinesin motor domain
binds to 13-tubulin and decorates microtubules with a B-surface
lattice. Proc Nat/ Acad Sci USA 1993, 90:1671-1675.
Decoration of microtubules with recombinant kinesin head domains
shows that microtubules have a B-type lattice of dimers. The stoichiometry
is one kinesin head per tubulin dimer, and chemical cross-linking connects kinesin to [[]-tubulin.
13.
•
Harrison B, Marchese-Ragona S, Gilbert S, Cheng N, Steven
A, Johnson KA: Decoration of the microtubule surface by one
kinesin head per tubulin heterodimer. Nature 1993, 362:73-7,5.
Describes microtubules decorated with kinesin head domains and imaged by cryo-electron microscopy.
S o n g Y-H, Mandelkow E: Anatomy of flagellar microtubules:
polarity, seam, junctions, and lattice. J Cell Biol 1995,
128:81-94.
Analysis of flagellar outer doublet and central pair microtubules by decoration with kinesin heads, showing the lattices (B-type in all cases) and
the polarity of dimers (u.-tubulin at the plus end).
14.
•
15.
Mitchison T): Localization of an exchangeable GTP-binding site
•
at the plus end of microtubules. Science 1993, 261:1044-1047.
A direct demonstration that the exchangeable GTP on 13-tubulin is near
the plus end of microtubules.
16.
Kuo SC, Sheetz MP: Force of single kinesin molecules meas•
ured with optical tweezers. Science 1993, 260:232-234.
This series of papers [16*-20 "] provides examples of how the micromechanics of molecular motors can be analyzed by laser tweezers, video
microscopy, and related techniques. The kinesin step sizes are related to
the tubulin dimer repeat, and the forces are in the picoNewton range per
kinesin head.
Svoboda K, Schmidt CF, Schnapp BJ, Block SM: Direct observation of kinesin stepping by optical trapping interferomelry.
Nature 1993, 365:721-727.
Demonstration by optical trapping that kinesin moves along microtubules
with step sizes of 8 nm. See also [16"] annotation.
17.
•
18.
Svoboda K, Block SM: Force and velocity measured for single
•
kinesin molecules. Cell 1994, 77:773-784.
Force-velocity relationships of kinesin coupled to silica beads and moving along microtubules argue that ATP hydrolysis and kinesin displacement are only loosely coupled. See also [16*] annotation.
Hunt AJ, Gittes F, Howard J: The force exerted by a single
kinesin molecule against a viscous load. Biophys J 1994,
67:766-781.
The speed of microtubules moving on a kinesin-coated surface was
measured in solvents of different viscosities. The results are compared
to different 'ratchet' or 'power stroke' models. See also [16 ° ] annotation.
19.
•
Hall K, Cole DG, Yeh Y, Scholey JM, Baskin RJ: Forcevelocity relationships in kinesin-driven motility. Nature 1993,
364:457-459.
A centrifuge microscope is used to measure force-velocity relationships
for kinesin, yielding force values in the range of 0.1 pN per kinesin head.
See also [16 •] annotation.
20.
*
21.
Ray S, Meyhofer E, Milligan RA, Howard J: Kinesin follows the microtubules protofilament axis. J Cell Biol 1993,
121:1083-1093.
22.
•
Hackney DD: Evidence for alternating head catalysis by kinesin
during microtubule-stimulated ATP hydrolysis. Proc Nat/Acad
Sci USA 1994, 91:6865~869.
StewartRJ, Thaler JP, Goldstein LSB: Direction of microtubule
movement is an intrinsic property of the motor domains of
kinesin heavy chain and Drosophila ncd protein. Proc Natl
Acad Sci USA 1993, 90:5209-5213.
The paper shows that the directionality of the movement is encoded in
the head domain of a motor, independently of its arrangement in the
protein.
27.
•
28.
Endow S, Kang S, Satterwhite L, Rose M, Skeen V, Salmon ED:
Yeast Kar3 is a minus-end microtubule motor protein that
destabilizes microtubules preferentially at the minus ends.
EMBO J 1994, 13:2708-2713.
29.
Middleton K, Carbon J: Kar3-encoded kinesin is a minus-enddirected motor that functions with centromere-binding proteins (cbf3) on an in-vitro yeast kinetochore. Proc Natl Acad
Sci USA 1994, 91:7212-7216.
30.
Hunt AJ, Howard J: Kinesin swivels to permit microtubule
movement in any direction. Proc Natl Acad Sci USA 1994,
90:11653-11657.
31.
BernsteinM, Beech PL, Katz SG, Rosenbaum JL: A new kinesinlike protein (kip1) localized to a single microtubule of the
Chlamydomonas flagellum. J Cell Biol 1994, 125:1313-1326.
Liao H, Li G, Yen TJ: Mitotic regulation of microtubule crosslinking activity of CENP-E kinetochore protein. Science 1994,
265:394-398.
An example of how a kinesin-like head domain can cooperate with
another microtubule-binding domain on the same protein under the
control of phosphorylation.
32.
•
33.
Sackett DL: Podophyllotoxin, steganacin and combretastatin:
natural products thai bind at the colchicine site of lubulin.
Pharmacol Ther 1993, 59:163-228.
34.
Hamel E: Interactions of tubulin with small ligands. In Microtubule proteins. Edited by Avila J. Boca Raton: CRC Press;
1990:89-191.
35.
Toso RJ, Jordan MA, Farrell KW, Matsumoto B, Wilson L: Kinetic stabilization of microtubule dynamic instability in vitro
by vinblastine. Biochemistry 1993, 32:1285-1293.
36.
Vandecandelaere A, Martin SR, Schilstra MJ, Bayley PM: Effects
of the tubulin-colchicine complex on microtubule dynamic instability. Biochemistry 1994, 33:2792-2801.
37.
Perez-RamirezB, Shearwin KE, Timasheff SN: The colchicineinduced GTPase activity of tubulin: state of the product activation by microtubule-promoting cosolvents. Biochemistry 1994,
33:6253-6261.
38.
Uppuluri S, Knipling L, Sackett DL, Wolff J: Localization of
the colchicine-binding site of tubulin. Proc Nail Acad Sci USA
1993, 90:11598-11602.
39.
Combeau C, Commercon A, Mioskowski C, Rousseau B,
Aubert F, Goeldner M: Predominant labeling of ~-tubulin
over ct-tubulin from porcine brain by a photoactivatable taxoid
derivative. Biochemistry 1994, 33:6676-6683.
40.
Rao S, Krauss N, Heerding J, Swindell C, Ringel I, Orr G,
Horwitz S: 3'-(p-azidobenzamido)taxol photolabels the Nterminal 31 amino-acids of ~-tubulin. J Biol Chem 1994,
269:3132-3134.
Microtubules and microtubule-associated proteins M a n d e l k o w and M a n d e l k o w
41.
Diaz JF, Andreu JM: Assembly of purified GDP tubulin into
microtubules induced by taxol and taxotere: reversibility,
ligand stoichiometry, and competition. Biochemistry 1993,
32:2747-2755.
This study distinguishes between the nucleation of microtubules on other
microtubules (template elongation) and on the pericentriolar material (astral nucleation) in Xenopus eggs, and describes the role of y-tubulin in
the nucleation activity.
42.
Shivanna BE), Mejillano MR, Williams TD, Himes RH: Exchangeable GTP binding-site of [3-tubulin: identification of
cysleine-12 as the major site of cross-linking by direct
pholoaffinity-labeling. J Bio/ Chem 1993, 268:127-132.
57.
•
43.
Jayaram B, Haley BE: Identification of peptides within the base
binding domains of the GTP-specific and ATP-specific bindingsites of tubulin. J Biol Chem 1994, 269:3233-3242.
Davis A, Sage CR, Dougherty CA, Farrell KW: Microtubule
dynamics modulated by guanosine triphosphate hydrolysis activity of ~-tubulin. Science 1994, 264:839-842.
This paper and [45"] provide examples of how yeast genetics can be
used to answer questions about tubulin structure, functional domains,
and GTPase activity.
44.
•
Reijo RA, Cooper EM, Beagle GJ, Huffaker TC: Systematic mutational analysis of the yeast [3-tubulin gene. Mol Biol Cell
1994, 5:2943.
A systematic mutation analysis of [~-tubulin (converting charged residues
to alanine) to reveal regions important for the functions of microtubules
in yeast.
45.
•
46.
47.
Rudiger M, Weber K: Characterization of the post-translational
modifications in tubulin from the marginal band of avian erythrocytes. Eur J Biochem 1993, 218:107-116.
Lu Q, Luduena RF: In vitro analysis of microtubule assembly
of isotypically pure tubulin dimers: intrinsic differences in the
assembly properties of c~-[~-ii, c~-13-iii,and (~-[3-iv tubulin dimers
in the absence of microtubule-associated protein. J Biol Chem
1994, 269:2041-2047.
48.
Ersfeld K, WehJand J, Plessmann U, Dodemont H, Gerke V,
Weber K: Characterization of the tubulin tyrosine ligase. J Cell
Biol 1993, 120:725-732.
49.
Paturle-Lafanechere L, Manier M, Trigault N, Pirollet F, Mazargull H, Job D: Accumulation of delta-2-tubulin, a major tubulin
variant that cannot be tyrosinated in neuronal tissues and in
stable microtubule assemblies. J Cell 5ci 1994, 107:1529-1543.
50.
Audebert S, Koulakoff A, Berwald-Netter Y, Gros F, Denoulet
P, Edde B: Developmental regulation of polyglutamylated c~tubulin and ~-tubulin in mouse-brain neurons. J Cell 5ci 1994,
107:2313-2322.
51.
Gurland G, Gundersen GG: Protein phosphatase inhibitors
induce the selective breakdown of stable microtubules in
fibroblasts and epithelial-cells. Proc Natl Acad 5ci USA 1993,
90:8827-8831
52.
Bachurski C, Theodorakis N, Coulson RM, Cleveland DW: An
amino-terminal tetrapeptide specifies cotranslational degradation of [3-tubulin but not ct-tubulin messenger RNAs. Mol Cell
Biol 1994, 14:4076~.086.
53.
Bramhill D, Thompson CM: GTP-dependent polymerization of
Escherichia coil FtsZ protein to form tubules. Proc Natl Acad
Sci USA ]994, 91:5813-5817.
Marchesi VT, Ngo N: In vitro assembly of multiprotein complexes containing cc-tubulin, [3-tubulin, and y-tubulin, heatshock protein hsp70, and elongation factor-lct. Proc Nat/Acad
Sci USA 1993, 90:3028-3032.
Protein compexes that represent centrosome precursors are isolated from
Chinese hamster ovary cells. They contain cytoskeletal proteins (a.-, [3-,
y-tubulin and actin) as well as elongation factors and chaperones.
54.
.
StearnsT, Kirschner M: In vitro reconstitution of centrosome
assemblyand function: the central role of y-tubulin. Cell 1994,
76:623-637.
A 25 S complex containing y-tubulin in Xenopus eggs can combine with
the sperm centriole to form the centrosome; this assembly requires ATP
but not microtubules. The complex confers the microtubule-nucleating
activity upon the centrosome.
55.
•
56.
•
Felix MA, Antony C, Wright M, Maro B: Centrosome assembly
in vitro: role of y-tubulin recruitment in Xenopus sperm aster
formation. J Biol Chem 1994, 124:19-31
Ahmad FJ, Joshi HC, Centonze VE, Baas PW: Inhibition of microtubule nucleation at the neuronal centrosome compromises
axon growth. Neuron 1994, 12:271-280.
Growth of axons depends on a supply of microtubules which are nucleated on the neuronal centrosomes and require the presence of ",f-tubulin.
Doxsey SJ, Stein P, Evans L, Calarco PD, Kirschner M: Pericentrin, a highly conserved centrosome protein involved in
microtubule organization. Cell 1994, 76:639-650.
Pericentrin is a newly discovered 220 kDa protein involved in organizing
the pericentriolar material and other structures from which microtubules
can be nucleated.
58.
•
59.
Maekawa T, Kuriyama R: Primary structure and microtubuleinteracting domain of the SP-H antigen: a mitotic MAP located
at the spindle pole and characterized as a homologous protein
to NuMa. J Cell Sci 1993, 105:589-600.
60.
Steffen W, Fajer EA, Linck RW: Centrosomal components
immunologically related to tektins from ciliary and flagellar
microtubules. J Cell 5ci 1994, 107:2095-2105.
61.
Horio T, Oakley BR: Human y-tubulin functions in fission yeast.
J Biol Chem 1994, 126:1465-1473.
Melki R, Vainberg IE, Chow RL, Cowan NJ: Chaperonin-mediated folding of vertebrate actin-related protein and y-tubulin.
J Cell Biol 1993, 122:1301-1310.
The actin homologue centractin and ~-tubulin, two centrosomal proteins,
are folded through the cytoplasmic chaperof~in complex TCP-1 ; the folding reaction requires ATP.
62.
•
63.
•
Sternlicht H, Farr GW, Sternlicht ML, Driscoll JK, Willison K,
Yaffe MB: The t-complex polypeptide-1 complex is a chaperonin for tubulin and actin in vivo. Proc Nat/ Acad Sci USA
1993, 90:9422-9426.
A pulse-labelling analysis of actin and tubulin biogenesis in Chinese
hamster ovary cells showing that thay must associate with the 900 kDa
TCP-1 complex in order to fold properly and associate into functional
tubulin heterodimers.
Gao YI, Vainberg IE, Chow RL, Cowan NJ: Two cofactors and
cytoplasmic chaperonin are required for the folding of cc-tubulin and [[3-tubulin. Mol Cell Biol 1993, 13:2478-2485.
Analysis of the interactions between the TCP-1 complex, actin, tubulin,
and other cofactors involved in the folding reaction.
64.
•
Melki R, Cowan NJ: Facilitated folding of actins and tubulins
occurs via a nucleotide-dependent interaction between cytoplasmic chaperonin and distinctive folding intermediates. Mol
Cell Biol 1994, 14:2895-2904.
This paper describes the sequence of steps in the folding of actin and
tubulin by the cytoplasmic chaperonin complex. The initial interaction
requires the chaperonin to be in the ADP-bound state, whereas release
requires ATP, which thus acts as a switching device.
65.
•
Chen XY, Sullivan DS, Huffaker TC: Two yeast genes with similarlty to TCP-1 are required for microtubule and actin function
in vivo, Proc Nat/ Acad 5ci USA 1994, 91:9111-9115.
The folding of actin and tubulin in yeast requires chaperonin complexes
similar to the TCP-1 complex.
66.
•
67.
Yu WQ, Baas PW: Changes in microtubule number and length
during axon differentiation. J Neurosci 1994, 14:2818-2829.
68.
Lo MMS, Fieles AW, Norris TE, Dargis PG, Caputo CB, Scott
CW, Lee VMY, Goedert M: Human tau isoforms confer distinct
morphological and functional properties to stably transfected
fibroblasts. Mol Brain Res 1993, 20:209-220.
69.
Degarcini EM, Delaluna S, Dominguez JE, Avila J: Overexpression of tau protein in COS-1 cells results in the stabilization of centrosome-independent microtubules and extension of
cytoplasmic processes. Mol Cell Biochem 1994, 130:187-196.
Kosik KS, McConlogue L: Microtubule-associated protein function: lessons from expression in Spodoptera frugiperda cells.
Cell Motil Cytoskeleton 1994, 28:195-198.
A review of how insect Sf9 cells can be used as a model system to generate 'neurite-like' cell processes by transfecting them with microtubuleassociated proteins.
70.
•
79
80
Cytoskeleton
71.
•
Baas PW, Pienkowski TP, Cimbalnik KA, Toyama K, Bakalis
S, Ahmad FJ, Kosik KS: Tau confers drug stability but not
cold stability to microtubules in living cells. J Cell Sci 1994,
107:135-143.
Expression of tau in insect Sf9 cells leads to the formation of 'neurite-like'
cell processes based on assembled microtubules. Probing their resistance
to depolymerizing drugs or low temperature shows that tau makes these
microtubules stable against drugs but not against cold.
87.
Bramblett GT, Goedert M, Jakes R, Merrick SE, Trojanowski
JQ, Lee VMY: Abnormal tau phosphorylation at Ser(396) in
Alzheimer's disease recapitulates development and contributes
to reduced microtubule binding. Neuron 1993, 10:1089-1099.
88.
Morishima-Kawashima M, Hasegawa M, Takio K, Suzuki M,
Titani K, Ihara Y: Ubiquitin is conjugated with amino-terminally processed tau in paired helical filaments. Neuron 1993,
10:1151-1160.
89.
Yan SD, Chen X, Schmidt A, Brett J, Godman G, Zou Y, Scott
CW, Caputo C, Frappier T, Smith MA et al.: Glycated tau
protein in Alzheimer-disease: a mechanism for induction of
oxidant stress. Proc Natl Acad Sci USA 1994, 91:7787-7791.
90.
Watanabe A, Hasegawa M, Suzuki M, Takio K, MorishimaKawashima M, Titani K, Arai T, Kosik KS, Ihara Y: In-vivo
phosphorylation sites in fetal and adult rat tau. J Biol Chem
1993, 268:257] 2-25717.
91.
Drewes G, Mandelkow E-M, Baumann K, Goris J, Merlevede W, Mandelkow E: Dephosphorylation of tau protein
and Alzheimer paired helical filaments by calcineurin and
phosphatase-2A. FEBS Lett 1993, 336:425-432.
92.
Schweers O, Sch(3nbrunn-Hanebeck E, Marx A, Mandelkow E:
Structural studies of tau protein and Alzheimer paired helical
filaments show no evidence for [~ structure. ] Biol Chem 1994,
269:24290-24297.
93.
wille H, Drewes G, Biernat J, Mandelkow E-M, Mandelkow E:
Alzheimer-like paired helical filaments and antiparallel dimers
formed from microtubule-associated protein tau in vitro. J Cell
Biol 1992, 118:573-584.
94.
Doll T, Meichsner M, Riederer BM, Honegger P, Matus A: An
isoform of mlcrotubule-associated proteln-2 (MAP2) containing 4 repeats of the tubulin-binding motif. J Cell Sci 1993,
106:633-640.
95.
Wallis K, Azhar S, Rho M, Lewis S, Cowan N, Murphy DB:
The mechanism of equilibrium binding of microtubule-associated protein-2 to microtubules: binding is a multi-phasic
process and exhibits positive cooperativity. J Biol Chem 1993,
268:15158-15167.
96.
Hagiwara H, Yorifuji H, Sato-Yoshitake R, Hirokawa N: Competition between motor molecules (kinesin and cytoplasmic
dynein) and fibrous microtubule-associated proteins in binding to mlcrotubules. J Biol Chem 1994, 269:3581-3589.
97.
Bassell GJ, Singer RH, Kosik KS: Association of poly(A)
messenger-RNA with microtubules in cultured neurons. Neuron 1994, 12:571-582.
Marya PK, Syed Z, Fraylich PE, Eagles PAM: Kinesin and
tau bind to distinct sties on microlubules. J Cell Sci 1994,
107:339-344.
98.
Georgieff I, Liem R, Couchie D, Mavilia C, Nunez J, Shelanski
M: Expression of high molecular weight tau in the central and
peripheral nervous systems. J Cell Sci 1993, 105:729-737.
Lopez LA, Sheetz MP: Steric inhibition of cytoplasmic dynein
and kinesin motility by MAP2. Cell Motil Cytoskeleton 1993,
24:1-16.
99.
Hammarback JA, Obar RA, Hughes SM, Vallee RB: MAPlb is
encoded as a polyprotein that is processed to form a complex N-terminal microtubule-binding domain. Neuron 1991,
7:129-139.
100.
Langkopf A, Hammarback J, Muller R, Vallee R, Garner
C: Microtubule-associated proteins 1A and LC2: two proteins encoded in one messenger RNA. J Biol Chem 1992,
267:16561 16566.
101.
Cravchik A, Reddy D, Matus A: Identification of a
novel microtubule-binding domain in microtubule-associated
protein-la (MAPla). J Cell Sci 1994, 107:661-672.
102.
Mann SS, Hammarback JA: Molecular characterization of light
chain-3, a microtubule-binding subunit of MAPla and MAPlb.
J Biol Chem 1994, 269:11492-11497.
103.
Chapin SJ, Bulinski JC: Cellular microtubules heterogeneous in
their content of microtubule-associated protein-4 (MAP4). Cell
Motil Cytoskeleton 1994, 27:133-149.
104.
Faruki S, Karsenti E: Purification of microtubule proteins from
Xenopus egg extracts: identification of a 230k MAP4-1ike protein. Cell Motil Cytoskeleton 1994, 28:108-118.
105.
Bokros C, Hugdahl J, Hanesworth V, Murthy J, Morejohn L:
Characterization of the reversible taxol-induced polymeriza-
72.
•
Barlow S, Gonzalez-Garay ML, West R, Olmsted JB, Cabral F:
Stable expression of heterologous microtubule-associated proteins (MAPs) in Chinese hamster ovary cells: evidence for differing roles of MAPs in microtubule organization. J Ce// Bio/
1994, 126:1017-1029.
COS cells were transfected with Drosophila 250 kDa MAP, human MAP4
and human tau. The exogenous MAPs had no major effect on the levels
of assembled microtubules or their drug sensitivity, suggesting that MAPs
may have roles distinct from those in microtubule stabilization.
73.
•
Harada A, Oguchi K, Okabe S, Kuno J, Terada S, Ohshima T,
Sato-Yoshitake R, Takei Y, Noda T, Hirokawa N: Altered microtubule organization in smaU-caliber axons of mice lacking
tau protein. Nature 1994, 369:488-49].
The first tau-null mouse appears to survive reasonably well, suggesting
that neurons have a back-up system for tau.
74.
Brandt R, Lee G: Functional organization of microtubule-associated protein tau: identification of regions which affect microtubule growth, nucleation, and bundle formation in vitro.
J Biol Chem 1993, 268:3414-3419.
75.
Brandt R, Lee G: Orientation, assembly, and stability of microtubule bundles induced by a fragment of tau protein. Cell
Motil Cytoskeleton 1994, 28:143-154.
76.
Gustke N, Trinczek B, Biemat J, Mandelkow E-M, Mandelkow
E: Domains of tau protein and interactions with microtubules.
Biochemistry ] 994, 33:95 ] 1-9522.
77.
Sanchez I, Cohen WD: Assembly and bundling of marginal
band microtubule protein: role of tau. Cell Motil Cytoskeleton
1994, 29:57-71.
78.
Goode BL, Feinstein SC: Identification of a novel microtubule
binding and assembly domain in the developmentally regulated
inter-repeat region of tau. J Cell Biol 1994, 124:769-782.
79.
80.
81.
82.
83.
Litman P, Barg J, Rindzoonski L, Ginzburg I: Subcellular localization of tau messenger RNA in differentiating neuronal
cell culture: implications for neuronal polarity. Neuron 1993,
10:627~38.
Wang Y, Loomis PA, Zinkowski RP, Binder LI: A novel tau
transcript in cultured human neuroblastoma cells expressing
nuclear tau. J Cell Biol 1993, 121:257-267.
Mandelkow E-M, Mandetkow E: Tau as a marker for
Alzheimer's disease. Trends Biochem Sci 1993, 18:480-483.
84,
•
Biernat J, Gustke N, Drewes G, Mandelkow E-M, Mandelkow E:
Phosphorylation of serine 262 strongly reduces the binding
of tau protein to microtubules: distinction between PHFlike immunoreactivity and microtubule binding. Neuron 1993,
11:153-]63.
This paper describes a critical residue in the repeat region of tau protein
whose phosphorylation detaches tau from microtubules and destabilizes
them.
85.
86.
Hasegawa M, Morishima-Kawashima M, Takio K, Suzuki M,
Titani K, Ihara Y: Protein sequence and mass spectrometric
analyses of tau in the Alzheimer's disease brain. J Biol Chem
1992, 26:1 7047-1 7054.
Yoshida H, Ihara Y: Tau in paired helical filaments is functionally distinct from fetal tau: assembly incompetence of paired
helical filament tau. J Neurochem 1993, 61:1183-1186.
M i c r o t u b u l e s and m i c r o t u b u l e - a s s o c i a t e d p r o t e i n s M a n d e l k o w and M a n d e l k o w
tion of plant tubulin into microtubules. Biochemistry 1993,
32:3437-3447.
118.
Cassimeris L: Regulation of microtubule dynamic instability.
Cell Motil Cytoskeleton 1993, 26:275-281.
106.
Hugdahl J, Bokrok C, Hanesworth V, Aalund G, Morejohn LC:
Unique functional characteristics of the polymerization and
MAP binding regulatory domains of plant tubulin. Plant Cell
1993, 5:1063-1080.
119.
Hush JM, Wadsworth P, Callaham DA, Hepler PK: Quantification of microtubule dynamics in living plant cells using fluorescence redistribution after photobleaching. J Cell Sci 1994,
107:775-784.
107.
Vantard M, Peter C, Fellous A, Schellenbaum P, Lambert
A: Characterization of a 100 kDal heat-stable microtubuleassociated protein from higher plants. Eur J Biochem 1994,
220:847-853.
120.
Lieuvin A, Labbe J-C, Dome M, Job D: Intrinsic microtubule
stability in interphase cells. J Cell Biol 1994, 124:985-996.
121.
Sawin KE, Mitchison TJ: Microtubule flux in mitosis is independent of chromosomes, centrosomes, and antiparallel microtubules. Mol Biol Cell 1994, 5:217-226.
122.
Zhai Y, Borisy GG: Quantitative determination of the
proportion of microtubule polymer present during the
mitosis-interphase transition. J Cell Sci 1994, 107:881-890.
123.
Mastronarde DN, McDonald KL, Ding R, Mclntosh JR: Interpolar spindle microtubules in PtK cells. J Cell Biol 1993,
123:1475-1489.
124.
Yuan M, Shaw PI, Warn RM, Lloyd CW: Dynamic reorientation
of cortical microtubules, from transverse to longitudinal, in living plant-cells. Proc Natl Acad Sci USA 1994, 91:6050-6053.
108.
109.
110.
111.
Keryer G, Luo ZJ, Cavadore JC, Erlichman J, Bornens M: Phosphorylation of the regulatory subunit of type-ii-~ cAMP-dependent protein kinase by cyclin B/p34(cdc2) kinase impairs its
binding to microtubule-associated protein-2. Proc Nat/ Acad
Sci USA 1993, 90:5418-5422.
Herskovits JS, Shpetner HS, Burgess CC, Vallee RB: Microtubules and src homology-3 domains stimulate the dynamin
GTPase via its C-terminal domain. Proc Natl Acad Sci USA
1993, 90:11468-11472.
Bollag G, McCormick F, Clark R: Characterization of full-length
neurofibromin: tubulin inhibits ras GAP activity. EMBO J 1993,
12:1923-1927.
125.
*
Roychowdhury S, Rasenick MM: Tubulin45 protein association
stabilizes GTP-binding and activates GTPase: cytoskeletal participation in neuronal signal transduction. Biochemistry 1994,
33:9800-9805.
126.
Rayment I, Holden HM, Whittaker M, Yohn CB, Lorenz M,
Holmes KC, Milligan RA: Structure of the actin-myosin complex and its implications for muscle contraction. Science 1993,
261:58-65.
127.
Tabony J: Morphological bifurcations involving reaction-diffusion processes during microtubule formation. Science 1994,
264:245-248.
An example of how tubulin assembly alone can lead to spatial and temporal order, independently of other cellular factors.
112.
Kirsch J, Betz H: Widespread expression of gephyrin, a putative
glycine receptor tubulin linker protein, in rat brain. Brain Res
1993, 621:301-310.
113.
Surridge CD, Burns RG: The difference in the binding of
phosphatidylino~itol distinguishes MAP2 from MAP2c and tau.
Biochemistry 1994, 33:8051-8057.
Schroder RR, Manstein DJ, Jahn W, Holden H, Rayment I,
Holmes KC, Spudich IA: Three-dimensional atomic model of
F-actin decorated with Dictyostelfum myosin-S1. Nature 1993,
364:171-174.
114.
*
McNally FJ, Vale RD: Identification of katanin, an ATPase
thai severs and disassembles stable microtubules. Cell 1993,
75:419-429.
Katanin is a heterodimeric protein complex that is capable of cutting
microtubules in the presence of ATP.
Wolf SG, Mosser G, Downing KH: Tubulin conformation
in zinc-induced sheets and macrotubes. J Struct Biol 1993,
111:190-199.
A high-resolution (4,~) electron-diffraction analysis of tubulin sheets, revealing the presence of secondary structure elements such as [8-sheets.
115.
•
129.
McLaughlin PJ, Gooch JT, Mannherz HG, Weeds AG: Structure
of gelsolin segment-l-actin complex and the mechanism of
filament severing. Nature 1993, 364:685-692.
130.
Noel J, Hamm H, Sigler P: The 2.2A crystal structure
of transducin-ct complexed with GTP-y-S. Nature 1993,
366:654-663.
131.
Wittinghofer A: The structure of transducin Gct: more to view
than just ras. Cell 1994, 76:201-204.
132.
Abrahams JP, Leslie A, Lutter R, Walker J: Structure at 2.8,~ resolution of F1-ATPase from bovine heart mitochondria. Nature
1994, 370:621-628.
Shiina N, Gotoh Y, Kubomura N, Iwamatsu A, Nishida E:
Microlubule severing by elongation factor 1-co. Science 1994,
266:282-285.
A new role - - severing of microtubules - - is discovered for the elongation factor-lu.. See also [116"].
116.
*
Durso NA, Cyr RJ: A calmodulin-sensitive interaction between
microtubules and a higher plant homolog of elongation factor
let. Plant Cell 1994, 6:893-905.
In this paper, elongation factor-l~ is shown to stabilize and bundle microtubules in a calmodulin-dependent fashion. See also [11S•].
117.
•
SandersMA, Salisbury JL: Centrin plays an essential role in microtubule severing during flagellar excision in Chlamydomonas
reinhardtii. J Biol Chem 1994, 124:795-805.
Severing of flagella in Chlamydomonas is achieved by cutting microtubules using contractile fibers that contain centrin, a calmodulin homoIogue.
128.
•
E Mandelkow and E-M Mandelkow, Max Planck Unit for Structural Molecular Biology, Notkestrasse 85, D-22603, Hamburg 52,
Germany.
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