Download Microtubule-based transport – basic

Commentary
ARTICLE SERIES: Cell Biology and Disease
2319
Microtubule-based transport – basic mechanisms,
traffic rules and role in neurological pathogenesis
Mariella A. M. Franker1 and Casper C. Hoogenraad1,2,*
1
Cell Biology, Faculty of Science, Utrecht University, 3584 CH Utrecht, The Netherlands
Department of Neuroscience, Erasmus Medical Center, 3000 DR Rotterdam, The Netherlands
2
*Author for correspondence ([email protected])
Journal of Cell Science 126, 2319–2329
ß 2013. Published by The Company of Biologists Ltd
doi: 10.1242/jcs.115030
Journal of Cell Science
Summary
Microtubule-based transport is essential for neuronal function because of the large distances that must be traveled by various building
blocks and cellular materials. Recent studies in various model systems have unraveled several regulatory mechanisms and traffic rules
that control the specificity, directionality and delivery of neuronal cargos. Local microtubule cues, opposing motor activity and cargoadaptors that regulate motor activity control microtubule-based transport in neurons. Impairment of intracellular transport is detrimental
to neurons and has emerged as a common factor in several neurological disorders. Genetic approaches have revealed strong links
between intracellular transport processes and the pathogenesis of neurological diseases in both the central and peripheral nervous
system. This Commentary highlights recent advances in these areas and discusses the transport defects that are associated with the
development of neurological diseases.
Key words: Brain, Neuron, Neurological disease, Microtubule, Transport, Kinesin, Dynein, Adaptor proteins
Introduction
To build and maintain highly complex neurons with active
synapses, many basic building blocks – for example, proteins,
lipids and cellular materials, such as mitochondria and synaptic
vesicles – need to be sorted to either axons or dendrites (Kapitein
and Hoogenraad, 2011; Namba et al., 2011; Rolls, 2011). It is
well known that intracellular cargo transport in neurons is driven
by motor proteins that can move directionally along either of two
types of cytoskeletal structures: actin filaments and microtubules
(see Box 1). Actin facilitates motility of motor proteins of the
myosin family, whereas microtubules serve as tracks for two
families of motor proteins, the kinesins and dyneins, which move
towards the microtubule plus-end or minus-end, respectively
(Hirokawa et al., 2009; Schliwa and Woehlke, 2003; Vale, 2003).
Evidence shows that microtubule-based transport mainly
facilitates the long-range transport into distal axons and
dendrites, whereas actin-based transport is important for shortrange transport and delivery of proteins to synapses and growth
cones (Dent et al., 2011; Hoogenraad and Bradke, 2009;
Kneussel and Wagner, 2013).
Although many members of the motor families participate in
neuronal cargo trafficking, the diversity of motor proteins alone is
insufficient to account for the specificity, directionality and timing
of delivery. It has been shown that regulation of transport occurs
through several additional processes, such as activation or
deactivation of motors, selective binding of motors to cargo,
through microtubule-associated proteins (MAPs) and microtubule
post-translational modifications (PTMs) (Akhmanova and
Hammer, 2010; Goldstein et al., 2008; Janke and Bulinski, 2011;
Schlager and Hoogenraad, 2009). How the microtubule-based
transport of specific cargo types is spatially and temporally
regulated in neurons is now an emerging field of investigation.
Neurons are especially vulnerable to defects in transport due to
the extreme length of the neuronal processes. The longest axon in
the human body can be up to 1 meter long, whereas an average
neuronal cell body is ,50 mm. Motor proteins and their regulators
have crucial roles in the development of many neurological and
neurodegenerative diseases (De Vos et al., 2008; Millecamps and
Julien, 2013). Disruption of intracellular transport can result in
vesicle-trafficking impairments, alter specific cargo interactions
and cause defects in retrograde survival signals, ultimately leading
to neuronal death and loss of brain function (Coleman, 2011;
Perlson et al., 2010; Saxena and Caroni, 2007). Axonal
pathologies, including abnormal accumulations of proteins and
organelles have been observed in patients with Huntington’s,
Parkinson’s and Alzheimer’s disease, and can severely hinder
neuronal transport pathways (Gunawardena and Goldstein,
2005; Millecamps and Julien, 2013). However, whether the
accumulations are the cause or the consequence of transport
defects is unclear, and the causal relationship between
transport impairments and neurodegeneration is unknown. The
identification of mutations in genes involved in neuronal transport
pathways strongly supports the view that defective transport can
directly trigger neurological diseases. Several disruptive mutations
in a- or b-tubulin and microtubule-based motors have been directly
linked to neurological diseases in both the central and peripheral
nervous system (Eschbach and Dupuis, 2011; Millecamps and
Julien, 2013; Tischfield et al., 2011). In this Commentary, we will
discuss the disease-causing mutations in motor and regulatory
proteins that lead to developmental and neurological disorders.
Before doing so, we outline the basic molecular mechanisms that
control neuronal transport and focus on microtubule cues,
bidirectional transport mechanisms and motor adaptors as
regulators of motor activity.
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Box 1. Biochemical and biophysical properties of
motor proteins
The movements of organelles were first observed in the squid
giant axon and, soon after, molecular motors were identified as the
effectors of this transport (Allen et al., 1982; Brady et al., 1982;
Vale et al., 1985). The decades following the discovery of motor
proteins have yielded many important insights into the molecular,
structural and biophysical properties of cytoskeleton-dependent
motor transport. Many motor properties, such as directionality, stall
forces, and step sizes are now well understood based on
controlled in vitro experiments (Block, 2007; Reck-Peterson et al.,
2006; Yildiz et al., 2008). The structure of kinesin was first
demonstrated using electron microscopy (Hirokawa et al., 2009),
and further high-resolution crystallographic structures of the motor
domains have led to detailed insights into how force and
movement are coupled (Carter et al., 2011; Kon et al., 2012;
Kull et al., 1996). Revolutionary single-molecule biophysics
experiments established that kinesin walks with an asymmetric
hand-over-hand stepping mechanism, hydrolyzing a single ATP
molecule per motor step (Asbury et al., 2003; Vale et al., 1996;
Yildiz et al., 2004). For the motor to be processive, there must be a
proper coordination between the two motor heads. It is still a
matter of debate how this coordination, or gating, takes place
(Block, 2007). Motor proteins move unidirectionally with either
plus-end motility (kinesins) or minus-end motility (dyneins).
However, occasional backstepping of motor proteins can occur
and was first observed for dynein (Toba et al., 2006). It has been
postulated that increased flexibility of the dynein structure gives
rise to a diffusional component in the dynein step, causing an
‘irregular’ stepping pattern that allows variable step sizes, back
steps and even sideways steps (DeWitt et al., 2012; Qiu et al.,
2012). Other studies show that kinesins are also capable of ATPdependent backstepping when they are biased by load (Carter and
Cross, 2005). In these situations, the neck-linker region plays an
important role in mediating strain between the two motor heads
and in regulating gating (Clancy et al., 2011).
Transport mechanisms in neurons
Microtubule-based transport is crucial for neuronal survival and
function, both during development and in the adult brain.
Different cellular processes, such as neuronal migration, axon
guidance, the establishment and maintenance of neuronal polarity
and synaptic plasticity, involve many cytoskeletal elements and
transport events that together form the complex cellular networks
of the nervous system and ensure that the proper connections are
made in the brain (Barnes and Polleux, 2009; Conde and Cáceres,
2009; Hoogenraad and Bradke, 2009). In this section, we will
briefly highlight the importance of the cytoskeletal network and
motor protein transport for neuronal development and function
(Fig. 1).
During development of the nervous system, neuron progenitor
cells migrate across the brain to their final destination (Hatten,
1999). Although extracellular factors and adhesion molecules
provide directional cues to direct the migrating cells and guide
axonal growth, cytoskeletal interactions produce the forces that
are necessary for nuclear positioning and neuronal migration
(Gundersen and Worman, 2013; Kuijpers and Hoogenraad, 2011;
Stiess and Bradke, 2011). For example, it has been shown that
actin-based myosin motors and the dynein regulator Lis1 (also
known as PAFAH1B1) are important for pulling the centrosome
and soma forwards during migration (Vallee et al., 2009). Most
neurons polarize during the migration process to form a distinct
axonal and somatodendritic compartment. It is well known that
microtubule-based mechanisms have an active role in determining
neuronal polarity and axonal outgrowth (Hoogenraad and
Bradke, 2009). For example, stable microtubules can act as
specific transport roads for the trafficking of signaling factors
that participate in neuronal polarization (Arimura and Kaibuchi,
2007; Witte and Bradke, 2008). After the polarization and
differentiation stages, neurons need to maintain their extensively
polarized morphology, and specific components must be
delivered to the correct neuronal compartment. Specific
targeting of different motor–cargo complexes to either the axon
or the dendrites ensures that the cargo is transported correctly
(Hirokawa et al., 2009; Kapitein and Hoogenraad, 2011). Recent
studies have found that, although a few kinesins are nonselective, most kinesin motors preferentially target cargos to the
axon (Huang and Banker, 2012) and only dynein transports cargo
exclusively into dendrites (Kapitein et al., 2010a). The
cytoskeleton organization in the axon initial segment (AIS) is
also thought to have a role in axonal targeting by preventing
vesicles with dendritic cargo from entering the axon (Song et al.,
2009; Watanabe et al., 2012). Moreover, mature neurons form
dendritic spines, which are small ‘knob’-shaped extensions
protruding from the surface of the dendrites. Dendritic spines
are the major post-synaptic sites for excitatory synaptic
transmission (Bourne and Harris, 2008; Kennedy et al., 2005).
Several studies have shown that dynamic microtubules
occasionally enter denditic spines and have a role in spine
morphology and synaptic plasticity (Hoogenraad and
Akhmanova, 2010). Microtubules in spines might influence
actin dynamics and promote the transport of postsynaptic cargos,
such as receptors and postsynaptic proteins. Moreover, transport
processes in the axon of neurons in the central and peripheral
nervous system are of key importance to distribute the various
building blocks and cellular materials (Hirokawa et al., 2010;
Millecamps and Julien, 2013; Perlson et al., 2010; Saxton and
Hollenbeck, 2012). Thus, many different microtubule-based
processes are actively involved in various phases of neuronal
development and in the mature nervous system.
Microtubule cues steer neuronal transport
Neuronal transport can be directly regulated by the underlying
microtubule cytoskeleton. The local organization of
microtubules, MAPs, such as axonal tau, and PTMs all affect
for neuronal transport (Janke and Kneussel, 2010; Verhey and
Hammond, 2009). In axons, anterograde and retrograde transport
is directly dictated by the unipolar organization of the
microtubule array (Hirokawa et al., 2009; Hoogenraad and
Bradke, 2009). However, in dendrites, the microtubules coalesce
into bundles of mixed polarity and a single motor type could
mediate bidirectional cargo transport by switching from
microtubules in which the minus-end is directed outwards
(minus-end out) to microtubules in which the plus-end is
directed outwards (plus-end out). The anti-parallel microtubule
organization in dendrites provides a structural framework for the
regulation of cargo trafficking and raises interesting models with
regard to neuronal trafficking rules (Kapitein and Hoogenraad,
2011; Rolls, 2011). Moreover, the influence of the local
organization of actin cytoskeleton at presynaptic terminals and
dendritic spines and its affect on neuronal cargo transport is of
significant importance (Hotulainen and Hoogenraad, 2010;
Transport and disease
A
B
Neuronal migration
Neuronal polarization
Synapse plasticity
Mixed microtubules
direct dendritic
transport
Dynein and Lis1
produce
pulling force
+ ++
Dynein
C
PSD
Postsynaptic
spine with
actin network
Actin maintains
spine shape
and scaffold
for the PSD
Dendrites
–
–– – Centrosome
–
–
+
+
++
Neuron protrusion
–
+
+
Microtubules
+
–
+
Nucleus
+
AIS filters
unwanted cargo
Actin
network
Postmitotic
neuron
Actin and myosin II
produce
contractile force
PSD
+
PTMs guide
axonal transport
Axon
Microtubules
grow inside
spines
–
Radial glial cell
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–
–
–
+
+
Microtubules
Microtubule entry
influences
synaptic activity
+
–
+
Key
Translocation
of nucleus
Direction of
migration
Fig. 1. The role of the cytoskeleton during neuronal development and maturation. (A) During maturation, neurons migrate to their final destination in the
brain, establish a highly polarized structure and form chemical synapses that make the connections between the pre- and post-synaptic sites. The cytoskeleton
plays crucial roles throughout these processes. During migration, actin-based myosin II motors and microtubule-based dynein motors generate the pulling and
pushing forces that translocate the nucleus (see also, Vallee et al., 2009). (B) To maintain the highly polarized neuronal structure, different cargos must be
selectively transported to either the somatodendritic or axonal compartment. The panel shows a few important mechanisms that guide neuronal polarity (see also,
Kapitein and Hoogenraad, 2011). Microtubule post-translational modification (PTMs) that are present in the axon (blue) but absent in dendrites (red) specifically
target axonal cargo, whereas microtubule bundles with mixed orientation direct transport in the dendrites. In addition, the axon initial segment (AIS) filters out
dendritic cargo and prevents it from entering the axon. (C) The actin network is the primary component that gives shape to the dendritic spines and forms a
scaffold for the post-synaptic density (PSD). In addition, microtubules have been shown to transiently enter spines (see Hoogenraad and Akhmanova, 2010).
Kneussel and Wagner, 2013). For example, myosin motors have
been shown to counteract microtubule-based transport and
facilitate the docking of mitochondria to actin filaments
(Saxton and Hollenbeck, 2012). In vitro approaches have
shown that the axonal MAP tau can be used to tune the
relative amounts of plus-end- and minus-end-directed kinesin
transport (Dixit et al., 2008; Vershinin et al., 2007). Low
concentrations of tau facilitate kinesin-mediated transport,
whereas high concentrations of tau, as is found in the distal
parts of the axon, are favorable for dynein-mediated transport.
Such a difference between the effects of tau on kinesin- and
dynein-based motility has not been confirmed in vivo, where
neither overexpression of tau nor tau deletion alters axonal
transport (Morfini et al., 2007; Yuan et al., 2008). Interestingly,
phosphorylated tau is involved in a number of neurological
disorders, such as Alzheimer’s disease, and has recently been
shown to mislocalize to dendritic spines and mediate synaptic
dysfunction (Hoover et al., 2010).
A leading model for driving selective neuronal transport is that
the affinity of motor proteins for microtubules is affected by the
PTMs on the microtubules (Janke and Kneussel, 2010; Verhey
and Hammond, 2009). For example, during neuronal
polarization, it is well known that vectorial membrane
trafficking occurs into the future axon (Bradke and Dotti,
1997), that stable microtubules act as specific transport roads
(Arimura and Kaibuchi, 2007; Witte and Bradke, 2008), and that
kinesin-1 motors accumulate specifically at the tips of axons
(Jacobson et al., 2006; Nakata and Hirokawa, 2003), all
suggesting that kinesin-1 prefers stable microtubules for axonal
cargo transport. Recent work has identified a specific region in
kinesin-1 that is responsible for its preference to bind to
detyrosinated microtubules (Konishi and Setou, 2009). Here,
knockdown of tubulin–tyrosin ligase (TTL) to decrease
tyrosinated microtubules resulted in the redistribution of
kinesin-1 in both axons and dendrites. Other data suggest that
the abundance of GTP-tubulin in microtubules at the AIS
underlies the selective kinesin-1 localization in axons (Nakata
et al., 2011). Therefore, selective transport is probably not
regulated by a single factor but probably by various local cues,
such as a combination of specific microtubule modifications,
associated proteins and tubulin conformations.
Opposing motor activity and bidirectional transport
Many and diverse types of proteins and cellular materials have
been shown to move bidirectionally in cells (Welte, 2004). A
large number of organelles and cytoskeleton proteins, including
endosomes (Deinhardt et al., 2006), autophagosomes (Maday
et al., 2012), intermediate filaments (Li et al., 2012) and synaptic
vesicles (Sabo and McAllister, 2003), are known to show
bidirectional movement in neuronal axons. Although there are
some differences between the movements of various neuronal
cargos, they share common features, such as frequent stopping
and pausing, as well as reversals in direction during transport,
and we are beginning to understand the importance of these
mechanisms to achieve appropriate cargo distributions in
neurons. An interesting example from Drosophila shows that
neuropeptide-containing dense core vesicles, which are
transported into the distal axons, initially pass the proximal
boutons, the presynaptic axonal bulges. Excess vesicles circulate
back to the soma and are occasionally captured in en passant
boutons, ensuring uniform distribution throughout the axon
(Wong et al., 2012).
Although bidirectional movement has been observed in almost
all cellular systems, it remains a matter of debate how these
processes are regulated. In vitro, motor proteins are intrinsically
capable of processively transporting cargo without requiring
additional regulation by other factors. As indicate above, a single
type of motor protein can mediate bidirectional movement on a
mixed network of opposite polarity microtubule bundles, such as
that found in the dendrites (Kapitein et al., 2010a). Another
interesting aspect of bidirectional transport is the interplay between
different microtubule motor types. Backwards and forwards
movements are typically observed during neuronal transport;
however, net movement in one direction occurs over large
distances in neurons. How is this achieved? One possibility is
that motors with opposite directionality are continuously acting on
a cargo and engage in a stochastic ‘tug-of-war’. This can result in
either stalling of the cargo or in bidirectional, salutatory movements
depending on the relative ‘strength’ of the attached motors.
Ultimately, one type of motor wins and transports the cargo in its
preferred direction. A recent study using synthetic ‘DNA origami’
found that ensembles of artificial cargo that are composed of
opposite polarity motors are engaged in a tug-of-war, resulting in
stalled cargo (Derr et al., 2012). Modeling has shown that stochastic
tug-of-war can induce salutatory bidirectional movements (Müller
et al., 2008), and deformations of endosomes during transport in
vivo in cohort with decreased speeds during transitions between
anterograde and retrograde transport (Soppina et al., 2009) argue in
favor of this model. In addition, purified neuronal vesicles that
move along microtubules in vitro show very similar behavior as
observed in vivo, suggesting that all components that are necessary
for transport are strongly attached to the vesicles themselves
(Hendricks et al., 2010). It is plausible, however, that motoradaptor proteins or other signaling proteins that are attached to the
vesicles were co-purified in this study and further regulated the
activity of the motors (Gross, 2003; Welte, 2004).
Thus, although there is evidence that supports a stochastic
‘tug-of-war’ hypothesis, the activity of opposing motors appears
to be closely interlinked (Fig. 2). Knockdown of either kinesin or
dynein results in impaired transport in both plus- and minus-end
directions in several different cell types (Jolly and Gelfand,
2011), and it has been shown that coupling an active plus-enddirected motor to peroxisomes also induces motility in the
opposite direction in Drosophila S2 cells (Ally et al., 2009)
suggesting that motors with opposite directionality directly
activate each other. Similar results were found in a study on
bidirectional movement of mitochondria in Drosophila larval
Kinases and
phosphatases
Rab GTPase
Ca2+
Cargo
Adaptor protein
Dynein
–
Journal of Cell Science
Journal of Cell Science 126 (11)
MAPs
Microtubule
Kinesin
PTMs
+
2322
Fig. 2. Bidirectional transport. Plus-end-directed kinesin motors and
minus-end-directed dynein motors are attached to the same cargo
simultaneously and can engage in a stochastic ‘tug-of-war’. In addition, these
motors are regulated further to ensure a proper distribution of cargo in living
cells (black arrows). Adaptor proteins mediate the specific binding between
motor proteins and their cargo. Crosstalk between adaptors and other
regulators is also possible, biasing movement in the direction of either
kinesin- (blue arrow) or dynein-mediated movement (green arrow). Several
signaling events, such as local changes in Ca2+ levels, phosphorylationdependent signaling and regulated Rab GTPase activity can activate or
deactivate motors or act on adaptor proteins and other regulators. Finally,
motor activity can be regulated by the underlying microtubule cytoskeleton,
including microtubule-associated proteins (MAPs) and microtubule posttranslational modifications (PTMs).
motor axons (Pilling et al., 2006). In contrast, several other
studies report that cargo transport in one direction is readily
achieved by selectively targeting a single type of motor protein,
and they demonstrate that the activation by a motor with an
opposite directionality is not required. For instance, a robust
unidirectional motility was observed when kinesin motors are
recruited to peroxisomes, even when dynein function was
disrupted (Kapitein et al., 2010b). Furthermore, acute inhibition
of either dynein heavy chain (DHC) or kinesin-1 heavy chain
(KHC, also known as KIF5B) with antibodies has been found
to only impair minus-end- and plus-end-directed motion,
respectively (Hendricks et al., 2010; Shubeita et al., 2008).
These apparently contradictory findings could reflect the various
control mechanisms that exist for motor-based protein transport
(Jolly and Gelfand, 2011). As will be discussed below in more
detail, coordination of opposing motors in neurons is a complex
process that is regulated by several molecular machineries at
various cellular levels.
Motor adaptors as regulators of motor activity
It is becoming increasingly clear that interactions between motors
and their adaptors are a crucial step in selective transport (Fig. 2)
(Akhmanova and Hammer, 2010; Goldstein et al., 2008;
Hirokawa et al., 2010; Schlager and Hoogenraad, 2009). A
recent report studying the axonal transport of vesicles containing
the prion protein (PrPC) showed that the composition of dynein
and kinesin motors is the same in anterograde-directed,
retrograde-directed and stationary vesicles (Encalada et al.,
2011). These data suggest that – in addition to local motor
recruitment mechanisms – a stable population of cargoassociated motors can be modulated by regulatory factors to
control anterograde and retrograde transport. Indeed,
microtubule-based transport is extensively controlled by
internal and external factors, and many regulators of dynein
and kinesin motors have been identified. Recent research has
Journal of Cell Science
Transport and disease
shown that adaptor proteins that link motors to their cargo can
function as either activators or inhibitors of motor proteins and
allow for changes in the direction of transport (Akhmanova and
Hammer, 2010; Schlager and Hoogenraad, 2009). The list of
molecules that are known to link transport motors to specific
neuronal cargos is rapidly expanding. Biochemical and
proteomics approaches, as well as high-throughput yeast twohybrid screens, have identified more than 100 proteins that are
associated with kinesin-1 in mammals, flies and worms
(Gindhart, 2006). Most of these proteins are cargo molecules
themselves, but some are adaptor proteins, including scaffolding
proteins and Rab GTPases that regulate neuronal transport. For
example, Jun N-terminal kinase (JNK)-interacting proteins (JIPs)
links kinesin-1 to cargo, such as reelin receptor apolipoprotein E
receptor 2 (ApoER2, also known as LRP8) and phosphorylated bamyloid precursor protein (APP), and MAP kinase-activating
death domain protein (MADD, also known as DENN or Rab3
GDP/GTP exchange factor) acts as an adaptor between kinesin-3
motors and synaptic vesicles (Hirokawa et al., 2010; Namba et al.,
2011; Schlager and Hoogenraad, 2009). Moreover, several
signaling events, such as local changes in Ca2+ levels,
phosphorylation-dependent signaling and regulated Rab GTPase
activity, have been found to influence neuronal transport and
regulate the loading and unloading of cargo (Fig. 2) (Namba
et al., 2011; Saxton and Hollenbeck, 2012; Schlager and
Hoogenraad, 2009). Here, we will give two examples of bidirectional cargo adaptors that control the balance between
dynein- and kinesin-based transport and regulate selective
transport in neurons.
Huntingtin, of which the mutated protein is implicated in
Huntingon’s disease, acts as a molecular switch between dynein
and kinesin-1 motors (Caviston et al., 2011; Colin et al., 2008).
When huntingtin is phosphorylated, kinesin is recruited to
vesicles, thereby biasing their movement along the microtubule
towards the plus-ends. In the unphosphorylated state, kinesin
recruitment to the vesicle is no longer stabilized, and dyneinmediated retrograde motility predominates (Colin et al., 2008).
Many other lines of research support the view that kinesin- and
dynein-mediated transport is modulated by phosphorylation. For
example, ERK1/2 phosphorylation of dynein intermediate chain
(DIC) recruits cytoplasmic dynein to signaling endosomes for
retrograde transport in axons (Mitchell et al., 2012). Moreover,
recent studies in Caenorhabditis elegans have shown that two
cyclin-dependent kinases, PCT-1 and CDK-5, directly polarize
trafficking of presynaptic components by inhibiting dyneinmediated retrograde transport and setting the balance between
anterograde and retrograde motors (Ou et al., 2010). In contrast,
in rat sensory neurons, CDK5 phosphorylation of the dyneininteracting protein NDEL1 stimulates the cargo transport
capacity of dynein (Pandey and Smith, 2011).
Adaptor proteins that interact with the mitochondrial outer
surface have been identified, and these are potential candidates to
regulate mitochondria transport throughout the neuron (Saxton
and Hollenbeck, 2012). Recent findings have demonstrated that
two very similar motor-adaptor proteins, TRAK1 and TRAK2
utilize different machineries to transport mitochondria to axons
and dendrites (van Spronsen et al., 2013). TRAK1 binds to both
kinesin-1 and dynein and directs mitochondria into axons,
whereas TRAK2 predominantly interacts with dynein–dynactin
and mediates the dendritic targeting of mitochondria. These
results are consistent with the polarized microtubule organization
2323
in neurons (Baas et al., 1988). Microtubules oriented with the
plus-end outwards drive the TRAK1–dynein–kinesin complex
into axons, whereas microtubules with mixed orientations allow
transport of the TRAK2–dynein complex into dendrites.
Interestingly, these data imply that the opposing motors are
closely interlinked in axons but can act independently from one
another in dendrites. The functional differences between TRAK1
and TRAK2 are explained by conformational differences; the
backfolding of TRAK2 affects its interaction with kinesin-1. It is
tempting to speculate that conformational switching of motor
proteins (see Box 2) and motor-adaptor proteins (as described
above) can coordinate bidirectional transport, polarized
trafficking and influence local cargo movements.
Box 2. Intramolecular motor–tail communication
Communication between the motor domain and tail region is an
important aspect of motor protein regulation. Recent evidence
suggests that the tail region of dynein, through its subunits or
associated proteins, controls its own motor activity. In vitro studies
found that the legs at odd angles (Loa) mutation F580Y in the tail
of the cytoplasmic dynein heavy chain (Hafezparast et al., 2003)
inhibits motor processivity (Ori-McKenney et al., 2010). In addition,
dynein-associated protein NudE (NDE1) and Lis1 are involved in
the interaction between the tail and motor domains of dynein (OriMcKenney et al., 2010; Zylkiewicz et al., 2011). NudE–Lis1mediated communication appears to be achieved through a
significantly different mechanism from the folding mechanism for
myosin V and some kinesins, which occurs through direct
intramolecular interactions between tail and motor domains (Adio
et al., 2006; Siththanandan and Sellers, 2011). Kinesin-1
undergoes a conformational change that brings the motor
domain in close contact with its tail region as shown by
sedimentation analysis (Hackney et al., 1992). This folded,
inactive conformation of kinesin-1 blocks processive motor
movement (Cai et al., 2007). The crystal structure of kinesin-1
and subsequent biochemical experiments suggest that steric
hindrance is not the cause of inhibition, but that instead the tail
region restrains ADP release (Kaan et al., 2011). Whereas binding
to cargo was initially proposed to unfold kinesin-1 and activate its
motility (Coy et al., 1999), it is now thought that motors can be
present in an inhibited state on the cargo surface and might
alternate between active and inactive conformations in response
to external signals (Blasius et al., 2007). In addition, singlemolecule in vitro studies using quantum dots have shown that
when kinesin-1 is in a folded inhibited state it is able to passively
diffuse along microtubules (Lu et al., 2009). It is plausible that the
ability to diffuse along microtubules is a general feature of
molecular motors. Similar mechanisms have been described for
the kinesin-5 family member Eg5 (also known as KIF11), which is
involved in positioning of the mitotic spindle (Kapitein et al., 2008),
for KIF1A, a kinesin-3 family member (Hammond et al., 2009),
and for MCAK, a kinesin-13 microtubule-depolymerizing motor
(Helenius et al., 2006). The motor properties described in Box 1,
such as motor back-stepping, motor-neck–tail communication and
diffusion might allow motors to slow down and step around
obstacles to resolve ‘road blocks’ or ‘traffic jams’, and to switch to
other cytoskeletal tracks without exchanging the motors attached
to the cargo. Indeed, in disease models of Alzheimer’s and motor
neuron disease, it appears that only a small fraction of cargos are
trapped in these ‘traffic jams’, suggesting that motors could
possibly bypass these blockages (Pilling et al., 2006; Shemesh
et al., 2008).
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Microtubule transport and disease
As microtubule-based transport is fundamental to the
development and function of mature neurons, defects in
neuronal transport pathways are likely to cause neuronal
diseases (Chevalier-Larsen and Holzbaur, 2006; De Vos et al.,
2008; Hirokawa et al., 2010; Salinas et al., 2008). Although many
studies suggest that transport pathways have a major role in the
onset and progression of various human developmental and
neurological disorders, the causal relationship between transport
impairments and neurodegeneration is largely unclear
(Gunawardena and Goldstein, 2005; Millecamps and Julien,
2013). Several kinesin-knockout mice display specific
neurological phenotypes, such as epilepsy and hydrocephalus
(Nakajima et al., 2012; Niwa et al., 2012), and kinesin-2-related
ciliopathy, causing hydrocephalus and disturbance in
dorsoventral patterning of the neural tube (Hirokawa et al.,
2012). The identification of mutations in genes involved in
neuronal transport pathways strengthens the direct link between
transport defects and human neurological diseases (Table 1 and
Fig. 3). In this section, we will discuss a few examples of
how mutations in different tubulin isotypes and motor proteins
might lead to defective neuronal transport and trigger
neurodegeneration.
One way to impair neuronal transport is to disrupt the
underlying microtubule network (Baas et al., 2005; Conde and
Cáceres, 2009; Janke and Bulinski, 2011; Lumb et al., 2012;
Tischfield et al., 2011). Mutations in a- and b-tubulin-encoding
genes have been identified in patients suffering from a wide
spectrum of neurological disorders, ranging from microcephaly
and seizures to intellectual disabilities and peripheral neuropathy.
For instance, mutations in the a-tubulin isotype VIII gene TUBA8
cause polymicrogyria with optic nerve hypoplasia (Abdollahi
et al., 2009), and mutations in the b-tubulin isotype II gene
TUBB2 result in asymmetrical polymicrogyria (Jaglin et al.,
2009). Moreover, in mice, mutations in the tubulin-specific
chaperone Tbce lead to a severe deficiency of microtubules in
distal axons in progressive motor neuronopathy (Martin et al.,
2002; Schaefer et al., 2007), and mutations in the microtubule
assembly and severing protein spastin 4 (SPG4) cause hereditary
spastic paraplegia in humans (Magariello et al., 2013; Nanetti
et al., 2012). In addition, missense mutations in TUBB3, which
encodes the neuron-specific b-tubulin isotype III, cause a more
Table 1. Mutations in transport proteins underlying neurological disorders
Mutant protein
Functional defect
Disease
Reference
Vesicular and organelle
transport
Defects in neuronal
migration; late-onset
motor neuron degeneration
Vesicular and organelle
transport
Synaptic vesicle transport,
mitochondrial transport
Axonal transport of K+dependent Na+/Ca2+
exchanger (NCKX2)
Impaired axonal transport
Lissencephaly; Charcot–Marie–
Tooth disease 2 (CMT2);
Perry syndrome; Spinal
muscular atropy
Hereditary spastic paraplegia
(SPG10); CMT2
CMT2
Congenital fibrosis of the
extraocular muscle (CFEOM)
(Braunstein et al., 2010; Harms
et al., 2010; Weedon et al.,
2011; Willemsen et al.,
2012)
(Crimella et al., 2012; Füger
et al., 2012)
(Gentil and Cooper, 2012;
Zhao et al., 2001)
(Desai et al., 2012; Lee et al.,
2012)
Impaired axonal transport;
axonal swelling; synapse
retraction
Impaired neuronal migration;
reduced axonal transport
Motor neuron disease;
amytrophic lateral sclerosis
(ALS); Perry syndrome
Lissencephaly
(Newsway et al., 2010;
Stockmann et al., 2012;
Uribe, 2010)
(Mokánszki et al., 2012)
a-tubulin isoform
Neuronal migration
Lissencephaly
TUBA8
a-tubulin isoform
Polymicrogyria with optic nerve
hypoplasia
TUBB2B
b-tubulin isoform
Local cell positioning
during cerebral cortical
development
Neuronal migration
(Cushion et al., 2013; Hikita
et al., 2013; Tischfield et al.,
2011)
(Abdollahi et al., 2009;
Tischfield et al., 2011)
TUBB3
b-tubulin isoform
(neuron specific)
Neuronal migrations and
axon guidance defects
TUBB5
TBCE
b-tubulin isoform
Tubulin chaperone
Neuronal migration
Impaired maintenance of
microtubules in motor
axons
Synaptic growth and
neurotransmission defects
CFEOM
Facial paralysis
Late-onset axonal
sensorimotor polyneuropathy
Microcephaly
Progressive motor
neuropathy
Mutations in motors
DYNC1H1 (dynein)
KIF5A
KIF1B
KIF21A
Motor adaptor and regulator
protein mutations
DCTN1/p150glued
(dynactin)
LIS1 (PAFAH1B1)
Cytoskeletal mutations
TUBA1A
SPG4 (spastin)
Functional role
Dynein regulation
Coupling of nucleus to
centrosome during
neuronal migration;
dynein regulator
Microtubule severer
Reduced transport of
synaptic vesicle proteins
Atrophy of ocular motor
muscles
Asymmetrical
polymicrogyria
Hereditary spastic paraplegia
(Cushion et al., 2013;
Romaniello et al., 2012;
Tischfield et al., 2011)
(Poirier et al., 2010; Tischfield
et al., 2011)
(Breuss et al., 2012)
(Martin et al., 2002; Schaefer
et al., 2007)
(Magariello et al., 2013;
Nanetti et al., 2012)
The table is limited to mutations in motor proteins and associated proteins that have a strong genetic link to neurological disease.
Transport and disease
2325
Tubulin
α-tubulin
β-tubulin
TUBB5
TUBB2B
TUBB3
Spastin
TUBA8
TUBA1A
Dendrite
–
+
Microtubule
TBCE
Microencephaly
Polymicrogyria
Lissencephaly
Congenital fibrosis of
the extraocular muscle
(CFEOM)
Motor neuropathy
Hereditary spastic
paraplegia (HSP)
Axon
Dendrite
Kinesin
Dynein
Mitochondrion
Amyotrohic lateral
sclerosis (ALS)
Cargo
Charcot–Marie–
Tooth 2 (CMT2)
Dynein
Dynactin
Lis1
+
–
Microtubule
KIF1B
–
+
Microtubule
Perry syndrome
Charcot–Marie–
Tooth 2 (CMT2)
Hereditary spastic
paraplegia (HSP)
Congenital fibrosis of
the extraocular muscle
(CFEOM)
Spinal muscular
atrophy (SMA)
KIF5A
KIF21A
–
Journal of Cell Science
Lissencenphaly
+
Microtubule
diffuse spectrum of brain malformations and neurological
disabilities, and certain phenotypes often segregate with
particular amino acid substitutions (Poirier et al., 2010;
Tischfield et al., 2010). A recent study directly investigated
whether TUBB3 mutations impair microtubule-based transport in
neurons and found that of the 14 b-tubulin mutants tested, only
two TUBB3 mutations directly disrupt the interaction with
kinesin motors and subsequently impair axonal transport (Niwa
et al., 2013). The other mutations in b-tubulin III probably disturb
other microtubule properties, such as the stability of a-tubulin–btubulin heterodimers or other characteristics, including the
(de)polymerization properties of tubulin, microtubule dynamics
or MAP interactions (Tischfield et al., 2011).
Several mutations in microtubule-based motors have also been
directly linked to neurological diseases. These motor mutations
can affect specific neuronal transport routes, or involve multiple
cargos and impair multiple pathways (Chevalier-Larsen and
Holzbaur, 2006; Hirokawa et al., 2010). Kinesin motor proteins
have been directly linked to several neurodegenerative diseases.
Mutations in the motor and stalk domain of the kinesin-1 family
member KIF5A cause hereditary spastic paraplegia and Charcot–
Marie–Tooth (CMT) disease type 2 (Crimella et al., 2012; Reid
et al., 2002). Mutations in the KIF21A stalk region have been
shown to be the underlying cause of congenital fibrosis of the
extraocular muscle type 1 (CFEOM1) (Yamada et al., 2003),
which is associated with defects in the oculomotor nerve and
results in an inability to move or raise the eyes. A recent study
Fig. 3. Major transport defects in
neurons and neurological diseases.
Impaired neuronal transport can be
caused by disruption of the microtubule
network, by changes in the processivity
or activation of motor proteins, or by
effects on the binding of cargo to motor
proteins. As illustrated here, interfering
with either of these regulatory
mechanisms gives rise to neuronal
transport defects and often leads to
variety of neurological disease
phenotypes. For example, mutations in
the kinesin motor protein KIF5A have
been linked to dominant forms of the
hereditary spastic paraplegia (HSP) and
Charcot–Marie–Tooth disease type 2
(CMT2) in humans, whereas mutations in
b-tubulin isotype III gene TUBB3 leads
to the ocular motility disorder, congenital
fibrosis of the extraocular muscles type 3
(CFEOM3). Furthermore, mutations in
dynein and its accessory proteins
dynactin and Lis1 have been associated
to spinal muscular atrophy (SMA),
amytrophic lateral sclerosis (ALS)-like
motor neuron degeneration and earlyonset parkinsonism called Perry
syndrome.
showed that knockdown of KIF21A causes the dysregulation of
Ca2+ at axonal boutons owing to inhibited transport of NCKX2, a
K+-dependent Na+-exchanger that is involved in Ca2+ clearance
in the large axon terminals of central neurons (Lee et al., 2012).
Moreover, a mutation in KIF1B-b, a kinesin-3 family member,
was identified in CMT2 patients (Zhao et al., 2001). The
heterogyzous Kif1b-knockout mouse presents similar phenotypes,
including a chronic peripheral neuropathy with progressive motor
weakness and motor coordination, and investigation of its sciatic
nerves revealed that transport of synaptic vesicles to the distal
axon is impaired. The KIF1B-b mutation is not the only gene that
is linked to CMT2, because, so far, mutations in more than 30
genes have been identified that cause CMT disease, several of
which are linked to specific transport pathways (Gentil and
Cooper, 2012).
Cytoplasmic dynein has a pivotal role in neuronal trafficking
because it is the only microtubule-based motor protein with
minus-end-directed motility (Pfister et al., 2006). Indeed,
mutations in the dynein heavy chain gene DHC1H1 lead to
striatal atrophy, neuronal migration defects, intellectual
disability, CMT and spinal muscular atrophy (SMA)
(Braunstein et al., 2010; Harms et al., 2010; Weedon et al.,
2011; Willemsen et al., 2012). Consistent with these findings,
mutations in dynein in Drosophila and C. elegans result in
intracellular protein aggregates and behavioral abnormalities,
including locomotor impairment (Eaton et al., 2002; Koushika
et al., 2004). Furthermore, disrupting the function of dynein in
Journal of Cell Science
2326
Journal of Cell Science 126 (11)
motor neurons in mice causes amyotrophic lateral sclerosis
(ALS)-like features (LaMonte et al., 2002; Teuling et al., 2008).
Mice strains with spontaneous heterozygous mutations in
DYNC1H1, such as the ‘legs at odd angles’ (Loa) and
‘cramping 1’ (Cra1) alleles, also show late-onset motor neuron
degeneration (Hafezparast et al., 2003. Recent in vitro studies
found that the Loa mutation F580Y in the tail of the mouse
cytoplasmic dynein heavy chain inhibits motor processivity (OriMcKenney et al., 2010). These data indicate that communication
between the motor domain and tail region is another important
aspect of motor regulation (see Box 2). Dynein is regulated by a
number of factors, such as LIS1, which, when disrupted, have
also been implicated in neurodevelopmental and neurological
diseases (Kardon and Vale, 2009). For example, missense
mutations in the dynactin gene (DCTN1) that encodes the
p150glued subunit of dynactin have been associated with both
familial and sporadic ALS (Münch et al., 2004; Puls et al., 2003;
Puls et al., 2005). Neuronal expression of mutant p150glued in
mice causes motor neuron disease, which is characterized by
defects in vesicular transport, axonal swelling and axo-terminal
degeneration (Chevalier-Larsen and Holzbaur, 2006; Lai et al.,
2007; Laird et al., 2008). It has recently been hypothesized that
the disease-causing mutations in the CAP-Gly domain of
p150glued inhibit the binding of dynactin at microtubule plusends, thereby abolishing the initiation of retrograde axonal
transport (Lloyd et al., 2012; Moughamian and Holzbaur, 2012).
Interestingly, different mutations in the same microtubulebinding domain of p150glued cause Perry syndrome, a
Parkinson-like disease that is characterized by the loss of
dopaminergic neurons (Farrer et al., 2009).
Another well-known neurological disorder that has been linked
to the dynein motor complex is lissencephaly, a developmental
brain malformation in which the surface of the brain is smooth
rather than full of folds and the disease subsequently leads to
seizures and mental retardation (Kato and Dobyns, 2003;
Kuijpers and Hoogenraad, 2011; Métin et al., 2008; WynshawBoris et al., 2010). Perturbing the levels of Lis1 causes defects in
microtubule-based processes, such as nuclear migration and
cargo transport (Pandey and Smith, 2011; Tanaka et al., 2004;
Tsai et al., 2007). Lis1 has been found to function in microtubulebased trafficking by preparing dynein-coupled cargos for
transport by targeting it to the plus-ends of microtubules
(Egan et al., 2012; Li et al., 2005; McKenney et al., 2010; Yi
et al., 2011). Recent single-molecule and structural studies
demonstrated that Lis1 binds in between the ATPase and
microtubule-binding domains of dynein and prolongs its
attachments to microtubules (Huang and Banker, 2012). Lis1
– at least in yeast – is not required for dynein-based cargo
motility once dynein is moving, but it instead has a general role
in initiating dynein-driven motility (Egan et al., 2012). The
numerous mutations in dynein, dynactin and associated
proteins, and the broad spectrum of phenotypes, demand a
clinical re-classification of dynein-related disorders, where the
neurological phenotype must be defined not only genetically
but also by the specific cellular effects. Future studies should
aim at dissecting the complexities of the dynein structure and
its interaction with associated proteins to find out the various
mechanisms by which disease-related dynein mutations disrupt
transport functions in the central and peripheral nervous
system.
Conclusions
Transport is essential for many different processes in neurons in
order to establish and maintain polarity, for axon guidance and
regeneration, as well as for synaptic plasticity, both during
developmental stages and in the adult brain. Although the
biochemical mechanisms underlying motor proteins-based
transport are well understood, much of the regulatory processes
that act on these motors are still unknown. Many studies now
show the importance of these regulatory pathways and emphasize
the role of adaptor proteins and regulators, in particular with
relevance to diseases of the brain. However, a number of
questions remain. How does a motor find the appropriate cargo?
How many different motor types are attached to a specific cargo?
How does the cargo know when to stop and ‘unload’ its content?
Disruption of transport is thought to be an early and perhaps
causative event in Alzheimer’s disease, Huntington’s disease and
ALS, in which abnormal accumulations of cytoskeleton proteins
and organelles in axons have been observed (De Vos et al., 2008;
Millecamps and Julien, 2013). The pivotal challenges are to
identify the molecular and cellular cause of human brain
disorders that arise from intracellular transport defects and to
determine how changes in motor protein regulation impact on
neuronal structure and function. Defects in transport lead to many
different and stress-related responses in cells, making it difficult
to elucidate the exact mechanisms underlying a disease. To
better understand intracellular transport in neurons, a better
understanding of the organization and dynamics of underlying
microtubule cytoskeleton, as well as novel approaches to control
and manipulate neuronal cargo transport are required (Jenkins
et al., 2012; Kapitein et al., 2010a). The spatial and temporal
arrangement of transport complexes and the cytoskeleton along
which they move is of crucial importance in order to understand
how multiple motors can regulate the transport and delivery of
specific cargos. These findings could potentially lead to new
tools that will be useful when investigating the transport
machinery in neurodegenerative disease models. Future studies
using genetic disease mouse models will advance our
understanding of the neurodegenerative disorders. Functional
assays in primary neuronal cells from mouse models or neurons
developed from inducible pluripotent stem (IPS) cells of patients
(Abeliovich and Doege, 2009) will further help to elucidate the
specific disease mechanisms. Moreover, advances in the
discovery and synthesis of small molecules that can modulate
microtubule-based processes in the brain might offer new
therapeutic paradigms to treat neurological defects and
intervene in neurodegenerative processes. The use of the
microtubule-stabilizing drug taxol to decrease fibrotic scar
formation and promote axon regeneration in rodents after
spinal cord injury (Ertürk et al., 2007; Hellal et al., 2011) is a
promising step in this direction, and other similar approaches
hopefully will follow.
Acknowledgements
We thank Lukas Kapitein and Phebe Wulf for critical reading of the
manuscript.
Funding
The work of our laboratories is supported by Het Prinses Beatrix
Spierfonds; the Netherlands Organization for Scientific Research
(NWO-ALW-VICI); the Netherlands Organization for Health
Research and Development (ZonMW-TOP); the European Science
Transport and disease
Foundation (EURYI); and the EMBO Young Investigators Program
(YIP).
Journal of Cell Science
References
Abdollahi, M. R., Morrison, E., Sirey, T., Molnar, Z., Hayward, B. E., Carr, I. M.,
Springell, K., Woods, C. G., Ahmed, M., Hattingh, L. et al. (2009). Mutation of the
variant alpha-tubulin TUBA8 results in polymicrogyria with optic nerve hypoplasia.
Am. J. Hum. Genet. 85, 737-744.
Abeliovich, A. and Doege, C. A. (2009). Reprogramming therapeutics: iPS cell
prospects for neurodegenerative disease. Neuron 61, 337-339.
Adio, S., Reth, J., Bathe, F. and Woehlke, G. (2006). Review: regulation mechanisms
of Kinesin-1. J. Muscle Res. Cell Motil. 27, 153-160.
Akhmanova, A. and Hammer, J. A., III (2010). Linking molecular motors to
membrane cargo. Curr. Opin. Cell Biol. 22, 479-487.
Allen, R. D., Metuzals, J., Tasaki, I., Brady, S. T. and Gilbert, S. P. (1982). Fast
axonal transport in squid giant axon. Science 218, 1127-1129.
Ally, S., Larson, A. G., Barlan, K., Rice, S. E. and Gelfand, V. I. (2009). Oppositepolarity motors activate one another to trigger cargo transport in live cells. J. Cell
Biol. 187, 1071-1082.
Arimura, N. and Kaibuchi, K. (2007). Neuronal polarity: from extracellular signals to
intracellular mechanisms. Nat. Rev. Neurosci. 8, 194-205.
Asbury, C. L., Fehr, A. N. and Block, S. M. (2003). Kinesin moves by an asymmetric
hand-over-hand mechanism. Science 302, 2130-2134.
Baas, P. W., Deitch, J. S., Black, M. M. and Banker, G. A. (1988). Polarity orientation
of microtubules in hippocampal neurons: uniformity in the axon and nonuniformity in
the dendrite. Proc. Natl. Acad. Sci. USA 85, 8335-8339.
Baas, P. W., Karabay, A. and Qiang, L. (2005). Microtubules cut and run. Trends Cell
Biol. 15, 518-524.
Barnes, A. P. and Polleux, F. (2009). Establishment of axon-dendrite polarity in
developing neurons. Annu. Rev. Neurosci. 32, 347-381.
Blasius, T. L., Cai, D., Jih, G. T., Toret, C. P. and Verhey, K. J. (2007). Two binding
partners cooperate to activate the molecular motor Kinesin-1. J. Cell Biol. 176, 11-17.
Block, S. M. (2007). Kinesin motor mechanics: binding, stepping, tracking, gating, and
limping. Biophys. J. 92, 2986-2995.
Bourne, J. N. and Harris, K. M. (2008). Balancing structure and function at
hippocampal dendritic spines. Annu. Rev. Neurosci. 31, 47-67.
Bradke, F. and Dotti, C. G. (1997). Neuronal polarity: vectorial cytoplasmic flow
precedes axon formation. Neuron 19, 1175-1186.
Brady, S. T., Lasek, R. J. and Allen, R. D. (1982). Fast axonal transport in extruded
axoplasm from squid giant axon. Science 218, 1129-1131.
Braunstein, K. E., Eschbach, J., Ròna-Vörös, K., Soylu, R., Mikrouli, E., Larmet,
Y., René, F., Gonzalez De Aguilar, J. L., Loeffler, J. P., Müller, H. P. et al. (2010).
A point mutation in the dynein heavy chain gene leads to striatal atrophy and
compromises neurite outgrowth of striatal neurons. Hum. Mol. Genet. 19, 4385-4398.
Breuss, M., Heng, J. I., Poirier, K., Tian, G., Jaglin, X. H., Qu, Z., Braun, A.,
Gstrein, T., Ngo, L., Haas, M. et al. (2012). Mutations in the b-tubulin gene TUBB5
cause microcephaly with structural brain abnormalities. Cell Rep. 2, 1554-1562.
Cai, D., Hoppe, A. D., Swanson, J. A. and Verhey, K. J. (2007). Kinesin-1 structural
organization and conformational changes revealed by FRET stoichiometry in live
cells. J. Cell Biol. 176, 51-63.
Carter, N. J. and Cross, R. A. (2005). Mechanics of the kinesin step. Nature 435, 308312.
Carter, A. P., Cho, C., Jin, L. and Vale, R. D. (2011). Crystal structure of the dynein
motor domain. Science 331, 1159-1165.
Caviston, J. P., Zajac, A. L., Tokito, M. and Holzbaur, E. L. (2011). Huntingtin
coordinates the dynein-mediated dynamic positioning of endosomes and lysosomes.
Mol. Biol. Cell 22, 478-492.
Chevalier-Larsen, E. and Holzbaur, E. L. (2006). Axonal transport and neurodegenerative
disease. Biochim. Biophys. Acta 1762, 1094-1108.
Clancy, B. E., Behnke-Parks, W. M., Andreasson, J. O., Rosenfeld, S. S. and Block,
S. M. (2011). A universal pathway for kinesin stepping. Nat. Struct. Mol. Biol. 18,
1020-1027.
Coleman, M. (2011). Molecular signaling how do axons die? Adv. Genet. 73, 185-217.
Colin, E., Zala, D., Liot, G., Rangone, H., Borrell-Pagès, M., Li, X. J., Saudou,
F. and Humbert, S. (2008). Huntingtin phosphorylation acts as a molecular switch
for anterograde/retrograde transport in neurons. EMBO J. 27, 2124-2134.
Conde, C. and Cáceres, A. (2009). Microtubule assembly, organization and dynamics
in axons and dendrites. Nat. Rev. Neurosci. 10, 319-332.
Coy, D. L., Hancock, W. O., Wagenbach, M. and Howard, J. (1999). Kinesin’s tail
domain is an inhibitory regulator of the motor domain. Nat. Cell Biol. 1, 288-292.
Crimella, C., Baschirotto, C., Arnoldi, A., Tonelli, A., Tenderini, E., Airoldi, G.,
Martinuzzi, A., Trabacca, A., Losito, L., Scarlato, M. et al. (2012). Mutations in
the motor and stalk domains of KIF5A in spastic paraplegia type 10 and in axonal
Charcot-Marie-Tooth type 2. Clin. Genet. 82, 157-164.
Cushion, T. D., Dobyns, W. B., Mullins, J. G., Stoodley, N., Chung, S. K., Fry, A. E.,
Hehr, U., Gunny, R., Aylsworth, A. S., Prabhakar, P. et al. (2013). Overlapping
cortical malformations and mutations in TUBB2B and TUBA1A. Brain 136, 536-548.
De Vos, K. J., Grierson, A. J., Ackerley, S. and Miller, C. C. (2008). Role of axonal
transport in neurodegenerative diseases. Annu. Rev. Neurosci. 31, 151-173.
Deinhardt, K., Salinas, S., Verastegui, C., Watson, R., Worth, D., Hanrahan, S.,
Bucci, C. and Schiavo, G. (2006). Rab5 and Rab7 control endocytic sorting along the
axonal retrograde transport pathway. Neuron 52, 293-305.
2327
Dent, E. W., Gupton, S. L. and Gertler, F. B. (2011). The growth cone cytoskeleton in
axon outgrowth and guidance. Cold Spring Harb. Perspect. Biol. 3, a001800.
Derr, N. D., Goodman, B. S., Jungmann, R., Leschziner, A. E., Shih, W. M. and
Reck-Peterson, S. L. (2012). Tug-of-war in motor protein ensembles revealed with a
programmable DNA origami scaffold. Science 338, 662-665.
Desai, J., Velo, M. P., Yamada, K., Overman, L. M. and Engle, E. C. (2012).
Spatiotemporal expression pattern of KIF21A during normal embryonic development
and in congenital fibrosis of the extraocular muscles type 1 (CFEOM1). Gene Expr.
Patterns 12, 180-188.
DeWitt, M. A., Chang, A. Y., Combs, P. A. and Yildiz, A. (2012). Cytoplasmic dynein
moves through uncoordinated stepping of the AAA+ ring domains. Science 335, 221225.
Dixit, R., Ross, J. L., Goldman, Y. E. and Holzbaur, E. L. (2008). Differential
regulation of dynein and kinesin motor proteins by tau. Science 319, 1086-1089.
Eaton, B. A., Fetter, R. D. and Davis, G. W. (2002). Dynactin is necessary for synapse
stabilization. Neuron 34, 729-741.
Egan, M. J., Tan, K. and Reck-Peterson, S. L. (2012). Lis1 is an initiation factor for
dynein-driven organelle transport. J. Cell Biol. 197, 971-982.
Encalada, S. E., Szpankowski, L., Xia, C. H. and Goldstein, L. S. (2011). Stable
kinesin and dynein assemblies drive the axonal transport of mammalian prion protein
vesicles. Cell 144, 551-565.
Ertürk, A., Hellal, F., Enes, J. and Bradke, F. (2007). Disorganized microtubules
underlie the formation of retraction bulbs and the failure of axonal regeneration.
J. Neurosci. 27, 9169-9180.
Eschbach, J. and Dupuis, L. (2011). Cytoplasmic dynein in neurodegeneration.
Pharmacol. Ther. 130, 348-363.
Farrer, M. J., Hulihan, M. M., Kachergus, J. M., Dächsel, J. C., Stoessl, A. J.,
Grantier, L. L., Calne, S., Calne, D. B., Lechevalier, B., Chapon, F. et al. (2009).
DCTN1 mutations in Perry syndrome. Nat. Genet. 41, 163-165.
Füger, P., Sreekumar, V., Schüle, R., Kern, J. V., Stanchev, D. T., Schneider, C. D.,
Karle, K. N., Daub, K. J., Siegert, V. K., Flötenmeyer, M. et al. (2012). Spastic
paraplegia mutation N256S in the neuronal microtubule motor KIF5A disrupts axonal
transport in a Drosophila HSP model. PLoS Genet. 8, e1003066.
Gentil, B. J. and Cooper, L. (2012). Molecular basis of axonal dysfunction and traffic
impairments in CMT. Brain Res. Bull. 88, 444-453.
Gindhart, J. G. (2006). Towards an understanding of kinesin-1 dependent transport
pathways through the study of protein-protein interactions. Brief Funct. Genomic
Proteomic 5, 74-86.
Goldstein, A. Y., Wang, X. and Schwarz, T. L. (2008). Axonal transport and the
delivery of pre-synaptic components. Curr. Opin. Neurobiol. 18, 495-503.
Gross, S. P. (2003). Dynactin: coordinating motors with opposite inclinations. Curr.
Biol. 13, R320-R322.
Gunawardena, S. and Goldstein, L. S. (2005). Polyglutamine diseases and transport
problems: deadly traffic jams on neuronal highways. Arch. Neurol. 62, 46-51.
Gundersen, G. G. and Worman, H. J. (2013). Nuclear positioning. Cell 152, 13761389.
Hackney, D. D., Levitt, J. D. and Suhan, J. (1992). Kinesin undergoes a 9 S to 6 S
conformational transition. J. Biol. Chem. 267, 8696-8701.
Hafezparast, M., Klocke, R., Ruhrberg, C., Marquardt, A., Ahmad-Annuar, A.,
Bowen, S., Lalli, G., Witherden, A. S., Hummerich, H., Nicholson, S. et al. (2003).
Mutations in dynein link motor neuron degeneration to defects in retrograde transport.
Science 300, 808-812.
Hammond, J. W., Cai, D., Blasius, T. L., Li, Z., Jiang, Y., Jih, G. T., Meyhofer,
E. and Verhey, K. J. (2009). Mammalian Kinesin-3 motors are dimeric in vivo and
move by processive motility upon release of autoinhibition. PLoS Biol. 7, e72.
Harms, M. B., Allred, P., Gardner, R., Jr, Fernandes Filho, J. A., Florence, J.,
Pestronk, A., Al-Lozi, M. and Baloh, R. H. (2010). Dominant spinal muscular
atrophy with lower extremity predominance: linkage to 14q32. Neurology 75, 539546.
Hatten, M. E. (1999). Central nervous system neuronal migration. Annu. Rev. Neurosci.
22, 511-539.
Helenius, J., Brouhard, G., Kalaidzidis, Y., Diez, S. and Howard, J. (2006). The
depolymerizing kinesin MCAK uses lattice diffusion to rapidly target microtubule
ends. Nature 441, 115-119.
Hellal, F., Hurtado, A., Ruschel, J., Flynn, K. C., Laskowski, C. J., Umlauf, M.,
Kapitein, L. C., Strikis, D., Lemmon, V., Bixby, J. et al. (2011). Microtubule
stabilization reduces scarring and causes axon regeneration after spinal cord injury.
Science 331, 928-931.
Hendricks, A. G., Perlson, E., Ross, J. L., Schroeder, H. W., III, Tokito, M. and
Holzbaur, E. L. (2010). Motor coordination via a tug-of-war mechanism drives
bidirectional vesicle transport. Curr. Biol. 20, 697-702.
Hikita, N., Hattori, H., Kato, M., Sakuma, S., Morotomi, Y., Ishida, H., Seto, T.,
Tanaka, K., Shimono, T., Shintaku, H. et al. (2013). A case of TUBA1A mutation
presenting with lissencephaly and Hirschsprung disease. Brain Dev.
Hirokawa, N., Noda, Y., Tanaka, Y. and Niwa, S. (2009). Kinesin superfamily motor
proteins and intracellular transport. Nat. Rev. Mol. Cell Biol. 10, 682-696.
Hirokawa, N., Niwa, S. and Tanaka, Y. (2010). Molecular motors in neurons: transport
mechanisms and roles in brain function, development, and disease. Neuron 68, 610638.
Hirokawa, N., Tanaka, Y. and Okada, Y. (2012). Cilia, KIF3 molecular motor and
nodal flow. Curr. Opin. Cell Biol. 24, 31-39.
Hoogenraad, C. C. and Akhmanova, A. (2010). Dendritic spine plasticity: new
regulatory roles of dynamic microtubules. Neuroscientist 16, 650-661.
Journal of Cell Science
2328
Journal of Cell Science 126 (11)
Hoogenraad, C. C. and Bradke, F. (2009). Control of neuronal polarity and
plasticity—a renaissance for microtubules? Trends Cell Biol. 19, 669-676.
Hoover, B. R., Reed, M. N., Su, J., Penrod, R. D., Kotilinek, L. A., Grant, M. K.,
Pitstick, R., Carlson, G. A., Lanier, L. M., Yuan, L. L. et al. (2010). Tau
mislocalization to dendritic spines mediates synaptic dysfunction independently of
neurodegeneration. Neuron 68, 1067-1081.
Hotulainen, P. and Hoogenraad, C. C. (2010). Actin in dendritic spines: connecting
dynamics to function. J. Cell Biol. 189, 619-629.
Huang, C. F. and Banker, G. (2012). The translocation selectivity of the kinesins that
mediate neuronal organelle transport. Traffic 13, 549-564.
Jacobson, C., Schnapp, B. and Banker, G. A. (2006). A change in the selective
translocation of the Kinesin-1 motor domain marks the initial specification of the
axon. Neuron 49, 797-804.
Jaglin, X. H., Poirier, K., Saillour, Y., Buhler, E., Tian, G., Bahi-Buisson, N., FalletBianco, C., Phan-Dinh-Tuy, F., Kong, X. P., Bomont, P. et al. (2009). Mutations in
the beta-tubulin gene TUBB2B result in asymmetrical polymicrogyria. Nat. Genet.
41, 746-752.
Janke, C. and Bulinski, J. C. (2011). Post-translational regulation of the microtubule
cytoskeleton: mechanisms and functions. Nat. Rev. Mol. Cell Biol. 12, 773-786.
Janke, C. and Kneussel, M. (2010). Tubulin post-translational modifications: encoding
functions on the neuronal microtubule cytoskeleton. Trends Neurosci. 33, 362-372.
Jenkins, B., Decker, H., Bentley, M., Luisi, J. and Banker, G. (2012). A novel split
kinesin assay identifies motor proteins that interact with distinct vesicle populations.
J. Cell Biol. 198, 749-761.
Jolly, A. L. and Gelfand, V. I. (2011). Bidirectional intracellular transport: utility and
mechanism. Biochem. Soc. Trans. 39, 1126-1130.
Kaan, H. Y., Hackney, D. D. and Kozielski, F. (2011). The structure of the kinesin-1
motor-tail complex reveals the mechanism of autoinhibition. Science 333, 883-885.
Kapitein, L. C. and Hoogenraad, C. C. (2011). Which way to go? Cytoskeletal
organization and polarized transport in neurons. Mol. Cell. Neurosci. 46, 9-20.
Kapitein, L. C., Kwok, B. H., Weinger, J. S., Schmidt, C. F., Kapoor, T. M. and
Peterman, E. J. (2008). Microtubule cross-linking triggers the directional motility of
kinesin-5. J. Cell Biol. 182, 421-428.
Kapitein, L. C., Schlager, M. A., Kuijpers, M., Wulf, P. S., van Spronsen, M.,
MacKintosh, F. C. and Hoogenraad, C. C. (2010a). Mixed microtubules steer
dynein-driven cargo transport into dendrites. Curr. Biol. 20, 290-299.
Kapitein, L. C., Schlager, M. A., van der Zwan, W. A., Wulf, P. S., Keijzer, N. and
Hoogenraad, C. C. (2010b). Probing intracellular motor protein activity using an
inducible cargo trafficking assay. Biophys. J. 99, 2143-2152.
Kardon, J. R. and Vale, R. D. (2009). Regulators of the cytoplasmic dynein motor. Nat.
Rev. Mol. Cell Biol. 10, 854-865.
Kato, M. and Dobyns, W. B. (2003). Lissencephaly and the molecular basis of neuronal
migration. Hum. Mol. Genet. 12 Spec No 1, R89-R96.
Kennedy, M. B., Beale, H. C., Carlisle, H. J. and Washburn, L. R. (2005). Integration
of biochemical signalling in spines. Nat. Rev. Neurosci. 6, 423-434.
Kneussel, M. and Wagner, W. (2013). Myosin motors at neuronal synapses: drivers of
membrane transport and actin dynamics. Nat. Rev. Neurosci. 14, 233-247.
Kon, T., Oyama, T., Shimo-Kon, R., Imamula, K., Shima, T., Sutoh, K. and Kurisu,
G. (2012). The 2.8 Å crystal structure of the dynein motor domain. Nature 484, 345350.
Konishi, Y. and Setou, M. (2009). Tubulin tyrosination navigates the kinesin-1 motor
domain to axons. Nat. Neurosci. 12, 559-567.
Koushika, S. P., Schaefer, A. M., Vincent, R., Willis, J. H., Bowerman, B. and
Nonet, M. L. (2004). Mutations in Caenorhabditis elegans cytoplasmic dynein
components reveal specificity of neuronal retrograde cargo. J. Neurosci. 24, 39073916.
Kuijpers, M. and Hoogenraad, C. C. (2011). Centrosomes, microtubules and neuronal
development. Mol. Cell. Neurosci. 48, 349-358.
Kull, F. J., Sablin, E. P., Lau, R., Fletterick, R. J. and Vale, R. D. (1996). Crystal
structure of the kinesin motor domain reveals a structural similarity to myosin. Nature
380, 550-555.
Lai, C., Lin, X., Chandran, J., Shim, H., Yang, W. J. and Cai, H. (2007). The G59S
mutation in p150(glued) causes dysfunction of dynactin in mice. J. Neurosci. 27,
13982-13990.
Laird, F. M., Farah, M. H., Ackerley, S., Hoke, A., Maragakis, N., Rothstein, J. D.,
Griffin, J., Price, D. L., Martin, L. J. and Wong, P. C. (2008). Motor neuron
disease occurring in a mutant dynactin mouse model is characterized by defects in
vesicular trafficking. J. Neurosci. 28, 1997-2005.
LaMonte, B. H., Wallace, K. E., Holloway, B. A., Shelly, S. S., Ascaño, J., Tokito,
M., Van Winkle, T., Howland, D. S. and Holzbaur, E. L. (2002). Disruption of
dynein/dynactin inhibits axonal transport in motor neurons causing late-onset
progressive degeneration. Neuron 34, 715-727.
Lee, K. H., Lee, J. S., Lee, D., Seog, D. H., Lytton, J., Ho, W. K. and Lee, S. H.
(2012). KIF21A-mediated axonal transport and selective endocytosis underlie the
polarized targeting of NCKX2. J. Neurosci. 32, 4102-4117.
Li, J., Lee, W. L. and Cooper, J. A. (2005). NudEL targets dynein to microtubule ends
through LIS1. Nat. Cell Biol. 7, 686-690.
Li, Y., Jung, P. and Brown, A. (2012). Axonal transport of neurofilaments: a single
population of intermittently moving polymers. J. Neurosci. 32, 746-758.
Lloyd, T. E., Machamer, J., O’Hara, K., Kim, J. H., Collins, S. E., Wong, M. Y.,
Sahin, B., Imlach, W., Yang, Y., Levitan, E. S. et al. (2012). The p150(Glued)
CAP-Gly domain regulates initiation of retrograde transport at synaptic termini.
Neuron 74, 344-360.
Lu, H., Ali, M. Y., Bookwalter, C. S., Warshaw, D. M. and Trybus, K. M. (2009).
Diffusive movement of processive kinesin-1 on microtubules. Traffic 10, 1429-1438.
Lumb, J. H., Connell, J. W., Allison, R. and Reid, E. (2012). The AAA ATPase
spastin links microtubule severing to membrane modelling. Biochim. Biophys. Acta
1823, 192-197.
Maday, S., Wallace, K. E. and Holzbaur, E. L. (2012). Autophagosomes initiate
distally and mature during transport toward the cell soma in primary neurons. J. Cell
Biol. 196, 407-417.
Magariello, A., Tortorella, C., Patitucci, A., Tortelli, R., Liguori, M., Mazzei, R.,
Conforti, F. L., Citrigno, L., Ungaro, C., Simone, I. L. et al. (2013). First mutation
in the nuclear localization signal sequence of spastin protein identified in a patient
with hereditary spastic paraplegia. Eur. J. Neurol. 20, e22-e23.
Martin, N., Jaubert, J., Gounon, P., Salido, E., Haase, G., Szatanik, M. and Guénet,
J. L. (2002). A missense mutation in Tbce causes progressive motor neuronopathy in
mice. Nat. Genet. 32, 443-447.
McKenney, R. J., Vershinin, M., Kunwar, A., Vallee, R. B. and Gross, S. P. (2010).
LIS1 and NudE induce a persistent dynein force-producing state. Cell 141, 304-314.
Métin, C., Vallee, R. B., Rakic, P. and Bhide, P. G. (2008). Modes and mishaps of
neuronal migration in the mammalian brain. J. Neurosci. 28, 11746-11752.
Millecamps, S. and Julien, J. P. (2013). Axonal transport deficits and neurodegenerative
diseases. Nat. Rev. Neurosci. 14, 161-176.
Mitchell, D. J., Blasier, K. R., Jeffery, E. D., Ross, M. W., Pullikuth, A. K., Suo, D.,
Park, J., Smiley, W. R., Lo, K. W., Shabanowitz, J. et al. (2012). Trk activation of
the ERK1/2 kinase pathway stimulates intermediate chain phosphorylation and
recruits cytoplasmic dynein to signaling endosomes for retrograde axonal transport.
J. Neurosci. 32, 15495-15510.
Mokánszki, A., Körhegyi, I., Szabó, N., Bereg, E., Gergev, G., Balogh, E., Bessenyei,
B., Sümegi, A., Morris-Rosendahl, D. J., Sztriha, L. et al. (2012). Lissencephaly
and band heterotopia: LIS1, TUBA1A, and DCX mutations in Hungary. J. Child
Neurol. 27, 1534-1540.
Morfini, G., Pigino, G., Mizuno, N., Kikkawa, M. and Brady, S. T. (2007). Tau
binding to microtubules does not directly affect microtubule-based vesicle motility.
J. Neurosci. Res. 85, 2620-2630.
Moughamian, A. J. and Holzbaur, E. L. (2012). Dynactin is required for transport
initiation from the distal axon. Neuron 74, 331-343.
Müller, M. J., Klumpp, S. and Lipowsky, R. (2008). Tug-of-war as a cooperative
mechanism for bidirectional cargo transport by molecular motors. Proc. Natl. Acad.
Sci. USA 105, 4609-4614.
Münch, C., Sedlmeier, R., Meyer, T., Homberg, V., Sperfeld, A. D., Kurt, A.,
Prudlo, J., Peraus, G., Hanemann, C. O., Stumm, G. et al. (2004). Point mutations
of the p150 subunit of dynactin (DCTN1) gene in ALS. Neurology 63, 724-726.
Nakajima, K., Yin, X., Takei, Y., Seog, D. H., Homma, N. and Hirokawa, N. (2012).
Molecular motor KIF5A is essential for GABA(A) receptor transport, and KIF5A
deletion causes epilepsy. Neuron 76, 945-961.
Nakata, T. and Hirokawa, N. (2003). Microtubules provide directional cues for
polarized axonal transport through interaction with kinesin motor head. J. Cell Biol.
162, 1045-1055.
Nakata, T., Niwa, S., Okada, Y., Perez, F. and Hirokawa, N. (2011). Preferential
binding of a kinesin-1 motor to GTP-tubulin-rich microtubules underlies polarized
vesicle transport. J. Cell Biol. 194, 245-255.
Namba, T., Nakamuta, S., Funahashi, Y. and Kaibuchi, K. (2011). The role of
selective transport in neuronal polarization. Dev. Neurobiol. 71, 445-457.
Nanetti, L., Baratta, S., Panzeri, M., Tomasello, C., Lovati, C., Azzollini, J., Gellera,
C., Di Bella, D., Taroni, F. and Mariotti, C. (2012). Novel and recurrent spastin
mutations in a large series of SPG4 Italian families. Neurosci. Lett. 528, 42-45.
Newsway, V., Fish, M., Rohrer, J. D., Majounie, E., Williams, N., Hack, M.,
Warren, J. D. and Morris, H. R. (2010). Perry syndrome due to the DCTN1 G71R
mutation: a distinctive levodopa responsive disorder with behavioral syndrome,
vertical gaze palsy, and respiratory failure. Mov. Disord. 25, 767-770.
Niwa, S., Nakajima, K., Miki, H., Minato, Y., Wang, D. and Hirokawa, N. (2012).
KIF19A is a microtubule-depolymerizing kinesin for ciliary length control. Dev. Cell
23, 1167-1175.
Niwa, S., Takahashi, H. and Hirokawa, N. (2013). b-Tubulin mutations that cause
severe neuropathies disrupt axonal transport. EMBO J.
Ori-McKenney, K. M., Xu, J., Gross, S. P. and Vallee, R. B. (2010). A cytoplasmic
dynein tail mutation impairs motor processivity. Nat. Cell Biol. 12, 1228-1234.
Ou, C. Y., Poon, V. Y., Maeder, C. I., Watanabe, S., Lehrman, E. K., Fu, A. K.,
Park, M., Fu, W. Y., Jorgensen, E. M., Ip, N. Y. et al. (2010). Two cyclindependent kinase pathways are essential for polarized trafficking of presynaptic
components. Cell 141, 846-858.
Pandey, J. P. and Smith, D. S. (2011). A Cdk5-dependent switch regulates Lis1/Ndel1/
dynein-driven organelle transport in adult axons. J. Neurosci. 31, 17207-17219.
Perlson, E., Maday, S., Fu, M. M., Moughamian, A. J. and Holzbaur, E. L. (2010).
Retrograde axonal transport: pathways to cell death? Trends Neurosci. 33, 335-344.
Pfister, K. K., Shah, P. R., Hummerich, H., Russ, A., Cotton, J., Annuar, A. A.,
King, S. M. and Fisher, E. M. (2006). Genetic analysis of the cytoplasmic dynein
subunit families. PLoS Genet. 2, e1.
Pilling, A. D., Horiuchi, D., Lively, C. M. and Saxton, W. M. (2006). Kinesin-1 and
Dynein are the primary motors for fast transport of mitochondria in Drosophila motor
axons. Mol. Biol. Cell 17, 2057-2068.
Poirier, K., Saillour, Y., Bahi-Buisson, N., Jaglin, X. H., Fallet-Bianco, C., Nabbout,
R., Castelnau-Ptakhine, L., Roubertie, A., Attie-Bitach, T., Desguerre, I. et al.
(2010). Mutations in the neuronal b-tubulin subunit TUBB3 result in malformation of
Journal of Cell Science
Transport and disease
cortical development and neuronal migration defects. Hum. Mol. Genet. 19, 44624473.
Puls, I., Jonnakuty, C., LaMonte, B. H., Holzbaur, E. L., Tokito, M., Mann, E.,
Floeter, M. K., Bidus, K., Drayna, D., Oh, S. J. et al. (2003). Mutant dynactin in
motor neuron disease. Nat. Genet. 33, 455-456.
Puls, I., Oh, S. J., Sumner, C. J., Wallace, K. E., Floeter, M. K., Mann, E. A.,
Kennedy, W. R., Wendelschafer-Crabb, G., Vortmeyer, A., Powers, R. et al.
(2005). Distal spinal and bulbar muscular atrophy caused by dynactin mutation. Ann.
Neurol. 57, 687-694.
Qiu, W., Derr, N. D., Goodman, B. S., Villa, E., Wu, D., Shih, W. and ReckPeterson, S. L. (2012). Dynein achieves processive motion using both stochastic and
coordinated stepping. Nat. Struct. Mol. Biol. 19, 193-200.
Reck-Peterson, S. L., Yildiz, A., Carter, A. P., Gennerich, A., Zhang, N. and Vale,
R. D. (2006). Single-molecule analysis of dynein processivity and stepping behavior.
Cell 126, 335-348.
Reid, E., Kloos, M., Ashley-Koch, A., Hughes, L., Bevan, S., Svenson, I. K.,
Graham, F. L., Gaskell, P. C., Dearlove, A., Pericak-Vance, M. A. et al. (2002). A
kinesin heavy chain (KIF5A) mutation in hereditary spastic paraplegia (SPG10). Am.
J. Hum. Genet. 71, 1189-1194.
Rolls, M. M. (2011). Neuronal polarity in Drosophila: sorting out axons and dendrites.
Dev. Neurobiol. 71, 419-429.
Romaniello, R., Tonelli, A., Arrigoni, F., Baschirotto, C., Triulzi, F., Bresolin, N.,
Bassi, M. T. and Borgatti, R. (2012). A novel mutation in the b-tubulin gene
TUBB2B associated with complex malformation of cortical development and deficits
in axonal guidance. Dev. Med. Child Neurol. 54, 765-769.
Sabo, S. L. and McAllister, A. K. (2003). Mobility and cycling of synaptic proteincontaining vesicles in axonal growth cone filopodia. Nat. Neurosci. 6, 1264-1269.
Salinas, S., Bilsland, L. G. and Schiavo, G. (2008). Molecular landmarks along the
axonal route: axonal transport in health and disease. Curr. Opin. Cell Biol. 20, 445453.
Saxena, S. and Caroni, P. (2007). Mechanisms of axon degeneration: from
development to disease. Prog. Neurobiol. 83, 174-191.
Saxton, W. M. and Hollenbeck, P. J. (2012). The axonal transport of mitochondria.
J. Cell Sci. 125, 2095-2104.
Schaefer, M. K., Schmalbruch, H., Buhler, E., Lopez, C., Martin, N., Guénet, J. L.
and Haase, G. (2007). Progressive motor neuronopathy: a critical role of the tubulin
chaperone TBCE in axonal tubulin routing from the Golgi apparatus. J. Neurosci. 27,
8779-8789.
Schlager, M. A. and Hoogenraad, C. C. (2009). Basic mechanisms for recognition and
transport of synaptic cargos. Mol. Brain 2, 25.
Schliwa, M. and Woehlke, G. (2003). Molecular motors. Nature 422, 759-765.
Shemesh, O. A., Erez, H., Ginzburg, I. and Spira, M. E. (2008). Tau-induced traffic
jams reflect organelles accumulation at points of microtubule polar mismatching.
Traffic 9, 458-471.
Shubeita, G. T., Tran, S. L., Xu, J., Vershinin, M., Cermelli, S., Cotton, S. L., Welte,
M. A. and Gross, S. P. (2008). Consequences of motor copy number on the
intracellular transport of kinesin-1-driven lipid droplets. Cell 135, 1098-1107.
Siththanandan, V. B. and Sellers, J. R. (2011). Regulation of myosin 5a and myosin
7a. Biochem. Soc. Trans. 39, 1136-1141.
Song, A. H., Wang, D., Chen, G., Li, Y., Luo, J., Duan, S. and Poo, M. M. (2009). A
selective filter for cytoplasmic transport at the axon initial segment. Cell 136, 11481160.
Soppina, V., Rai, A. K., Ramaiya, A. J., Barak, P. and Mallik, R. (2009). Tug-of-war
between dissimilar teams of microtubule motors regulates transport and fission of
endosomes. Proc. Natl. Acad. Sci. USA 106, 19381-19386.
Stiess, M. and Bradke, F. (2011). Neuronal polarization: the cytoskeleton leads the
way. Dev. Neurobiol. 71, 430-444.
Stockmann, M., Meyer-Ohlendorf, M., Achberger, K., Putz, S., Demestre, M., Yin,
H., Hendrich, C., Linta, L., Heinrich, J., Brunner, C. et al. (2012). The dynactin
p150 subunit: cell biology studies of sequence changes found in ALS/MND and
Parkinsonian Syndromes. J. Neural Transm. 120, 785-798.
Tanaka, T., Serneo, F. F., Higgins, C., Gambello, M. J., Wynshaw-Boris, A. and
Gleeson, J. G. (2004). Lis1 and doublecortin function with dynein to mediate
coupling of the nucleus to the centrosome in neuronal migration. J. Cell Biol. 165,
709-721.
Teuling, E., van Dis, V., Wulf, P. S., Haasdijk, E. D., Akhmanova, A., Hoogenraad,
C. C. and Jaarsma, D. (2008). A novel mouse model with impaired dynein/dynactin
function develops amyotrophic lateral sclerosis (ALS)-like features in motor neurons
and improves lifespan in SOD1-ALS mice. Hum. Mol. Genet. 17, 2849-2862.
Tischfield, M. A., Baris, H. N., Wu, C., Rudolph, G., Van Maldergem, L., He, W.,
Chan, W. M., Andrews, C., Demer, J. L., Robertson, R. L. et al. (2010). Human
TUBB3 mutations perturb microtubule dynamics, kinesin interactions, and axon
guidance. Cell 140, 74-87.
2329
Tischfield, M. A., Cederquist, G. Y., Gupta, M. L., Jr and Engle, E. C. (2011).
Phenotypic spectrum of the tubulin-related disorders and functional implications of
disease-causing mutations. Curr. Opin. Genet. Dev. 21, 286-294.
Toba, S., Watanabe, T. M., Yamaguchi-Okimoto, L., Toyoshima, Y. Y. and Higuchi,
H. (2006). Overlapping hand-over-hand mechanism of single molecular motility of
cytoplasmic dynein. Proc. Natl. Acad. Sci. USA 103, 5741-5745.
Tsai, J. W., Bremner, K. H. and Vallee, R. B. (2007). Dual subcellular roles for LIS1
and dynein in radial neuronal migration in live brain tissue. Nat. Neurosci. 10, 970979.
Uribe, V. (2010). DCTN1 mutations are implicated in multiple neurodegenerative
disorders. Clin. Genet. 77, 32-34.
Vale, R. D. (2003). The molecular motor toolbox for intracellular transport. Cell 112,
467-480.
Vale, R. D., Schnapp, B. J., Mitchison, T., Steuer, E., Reese, T. S. and Sheetz, M. P.
(1985). Different axoplasmic proteins generate movement in opposite directions along
microtubules in vitro. Cell 43, 623-632.
Vale, R. D., Funatsu, T., Pierce, D. W., Romberg, L., Harada, Y. and Yanagida,
T. (1996). Direct observation of single kinesin molecules moving along microtubules.
Nature 380, 451-453.
Vallee, R. B., Seale, G. E. and Tsai, J. W. (2009). Emerging roles for myosin II and
cytoplasmic dynein in migrating neurons and growth cones. Trends Cell Biol. 19, 347355.
van Spronsen, M., Mikhaylova, M., Lipka, J., Schlager, M. A., van den Heuvel,
D. J., Kuijpers, M., Wulf, P. S., Keijzer, N., Demmers, J., Kapitein, L. C. et al.
(2013). TRAK/Milton motor-adaptor proteins steer mitochondrial trafficking to axons
and dendrites. Neuron 77, 485-502.
Verhey, K. J. and Hammond, J. W. (2009). Traffic control: regulation of kinesin
motors. Nat. Rev. Mol. Cell Biol. 10, 765-777.
Vershinin, M., Carter, B. C., Razafsky, D. S., King, S. J. and Gross, S. P. (2007).
Multiple-motor based transport and its regulation by Tau. Proc. Natl. Acad. Sci. USA
104, 87-92.
Watanabe, K., Al-Bassam, S., Miyazaki, Y., Wandless, T. J., Webster, P. and
Arnold, D. B. (2012). Networks of polarized actin filaments in the axon initial
segment provide a mechanism for sorting axonal and dendritic proteins. Cell Rep. 2,
1546-1553.
Weedon, M. N., Hastings, R., Caswell, R., Xie, W., Paszkiewicz, K., Antoniadi, T.,
Williams, M., King, C., Greenhalgh, L., Newbury-Ecob, R. et al. (2011). Exome
sequencing identifies a DYNC1H1 mutation in a large pedigree with dominant axonal
Charcot-Marie-Tooth disease. Am. J. Hum. Genet. 89, 308-312.
Welte, M. A. (2004). Bidirectional transport along microtubules. Curr. Biol. 14, R525R537.
Willemsen, M. H., Vissers, L. E., Willemsen, M. A., van Bon, B. W., Kroes, T., de
Ligt, J., de Vries, B. B., Schoots, J., Lugtenberg, D., Hamel, B. C. et al. (2012).
Mutations in DYNC1H1 cause severe intellectual disability with neuronal migration
defects. J. Med. Genet. 49, 179-183.
Witte, H. and Bradke, F. (2008). The role of the cytoskeleton during neuronal
polarization. Curr. Opin. Neurobiol. 18, 479-487.
Wong, M. Y., Zhou, C., Shakiryanova, D., Lloyd, T. E., Deitcher, D. L. and Levitan,
E. S. (2012). Neuropeptide delivery to synapses by long-range vesicle circulation and
sporadic capture. Cell 148, 1029-1038.
Wynshaw-Boris, A., Pramparo, T., Youn, Y. H. and Hirotsune, S. (2010).
Lissencephaly: mechanistic insights from animal models and potential therapeutic
strategies. Semin. Cell Dev. Biol. 21, 823-830.
Yamada, K., Andrews, C., Chan, W. M., McKeown, C. A., Magli, A., de Berardinis,
T., Loewenstein, A., Lazar, M., O’Keefe, M., Letson, R. et al. (2003).
Heterozygous mutations of the kinesin KIF21A in congenital fibrosis of the
extraocular muscles type 1 (CFEOM1). Nat. Genet. 35, 318-321.
Yi, J. Y., Ori-McKenney, K. M., McKenney, R. J., Vershinin, M., Gross, S. P. and
Vallee, R. B. (2011). High-resolution imaging reveals indirect coordination of
opposite motors and a role for LIS1 in high-load axonal transport. J. Cell Biol. 195,
193-201.
Yildiz, A., Tomishige, M., Vale, R. D. and Selvin, P. R. (2004). Kinesin walks handover-hand. Science 303, 676-678.
Yildiz, A., Tomishige, M., Gennerich, A. and Vale, R. D. (2008). Intramolecular strain
coordinates kinesin stepping behavior along microtubules. Cell 134, 1030-1041.
Yuan, A., Kumar, A., Peterhoff, C., Duff, K. and Nixon, R. A. (2008). Axonal
transport rates in vivo are unaffected by tau deletion or overexpression in mice.
J. Neurosci. 28, 1682-1687.
Zhao, C., Takita, J., Tanaka, Y., Setou, M., Nakagawa, T., Takeda, S., Yang, H. W.,
Terada, S., Nakata, T., Takei, Y. et al. (2001). Charcot-Marie-Tooth disease type
2A caused by mutation in a microtubule motor KIF1Bbeta. Cell 105, 587-597.
Zyłkiewicz, E., Kijańska, M., Choi, W. C., Derewenda, U., Derewenda, Z. S. and
Stukenberg, P. T. (2011). The N-terminal coiled-coil of Ndel1 is a regulated scaffold
that recruits LIS1 to dynein. J. Cell Biol. 192, 433-445.