Download extracellular and intracellular signaling for neuronal polarity

Physiol Rev 95: 995–1024, 2015
Published July 1, 2015; doi:10.1152/physrev.00025.2014
EXTRACELLULAR AND INTRACELLULAR
SIGNALING FOR NEURONAL POLARITY
Takashi Namba, Yasuhiro Funahashi, Shinichi Nakamuta, Chundi Xu, Tetsuya Takano,
and Kozo Kaibuchi
Department of Cell Pharmacology, Nagoya University Graduate School of Medicine, Nagoya, Japan
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INTRODUCTION
DEVELOPMENT OF NEURONAL...
MECHANISMS OF NEURONAL...
THEORETICAL BASIS OF NEURONAL...
MAINTENANCE OF NEURONAL...
INVOLVEMENT OF NEURONAL...
NEW ERA OF NEURONAL POLARITY
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I. INTRODUCTION
The establishment of cell polarity is crucial for the development of tissues and organs (205, 311, 317). Various types of
cell polarity are found in the human body, including apicobasal polarity in epithelial cells, front-rear polarity in migrating cells, and axon-dendrite polarity in neurons. In general, neurons extend two distinct types of neurites: dendrites and axons. Axons and dendrites contain different
types of proteins and organelles and thus differ in both
function and morphology (FIGURE 1). Dendrites are relatively short and thick, and in most excitatory neurons, they
possess dendritic spines. Functionally, dendrites receive
chemical signals from other neurons through neurotransmitter receptors. Axons are typically long and thin, and
axon terminals contain synaptic vesicles for synaptic transmission to other neurons. Thus neurons are polarized both
functionally and morphologically (52). Many scientists are
fascinated by the fundamental issue in neuroscience: the
mechanisms of neuronal polarization. Although Santiago
Ramón y Cajal discovered the basic morphological features
of neurons using “la reazione near,” Golgi’s method, in the
early 20th century, there were few studies of neuronal po-
larization until the late 1980s (52). Dr. Banker and colleagues published the first report containing the phrase
“neuronal polarity” (41). These authors subsequently described the process of neuronal morphogenesis under culture conditions (66) and divided the developmental stages
of cultured hippocampal neurons according to their morphology (stages 1 to 5). Many researchers have attempted to
examine neuronal polarization based on these definitions.
In contrast to studies using cultured neurons, studies of
neuronal polarity in vivo remain under development. In
recent years, in utero electroporation methods for genetic
manipulation of the rodent brain have become feasible
(285, 328). This approach allows us to examine neuronal
polarity in the mouse cerebral cortex. In vivo, neuronal
polarity includes both axon-dendrite polarity and neuronal
migration.
Here, we review the current knowledge about neuronal polarization in vitro and in vivo, summarize the molecular
network regulating neuronal polarity, and discuss the future of research investigating neuronal polarity.
II. DEVELOPMENT OF NEURONAL
POLARIZATION IN VITRO AND IN VIVO
A. Neuronal Polarization in Cultured
Neurons
Cultured neurons dissociated from embryonic hippocampus have frequently been used to examine neuronal polar-
0031-9333/15 Copyright © 2015 the American Physiological Society
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Namba T, Funahashi Y, Nakamuta S, Xu C, Takano T, Kaibuchi K. Extracellular and
Intracellular Signaling for Neuronal Polarity. Physiol Rev 95: 995–1024, 2015; Published July 1, 2015; doi:10.1152/physrev.00025.2014.—Neurons are one of the
highly polarized cells in the body. One of the fundamental issues in neuroscience is how
neurons establish their polarity; therefore, this issue fascinates many scientists. Cultured neurons are useful tools for analyzing the mechanisms of neuronal polarization, and indeed,
most of the molecules important in their polarization were identified using culture systems. However,
we now know that the process of neuronal polarization in vivo differs in some respects from that in
cultured neurons. One of the major differences is their surrounding microenvironment; neurons in vivo
can be influenced by extrinsic factors from the microenvironment. Therefore, a major question remains: How are neurons polarized in vivo? Here, we begin by reviewing the process of neuronal
polarization in culture conditions and in vivo. We also survey the molecular mechanisms underlying
neuronal polarization. Finally, we introduce the theoretical basis of neuronal polarization and the
possible involvement of neuronal polarity in disease and traumatic brain injury.
NEURONAL POLARITY SIGNALING
ity. In that system, five developmental stages have been
defined (66) (FIGURE 1). First, hippocampal neurons form
several thin filopodia shortly after the dissociated neurons
are seeded (stage 1). Next, these neurons extend multiple
immature neurites, described as minor processes (stage 2).
These neurites undergo repeated random growth and retraction, and there are no morphological differences among
A
Cultured Neurons
Stage 1
Stage 2
Stage 3
Immature axon
C
Inheritance of polarity
(developing retina)
Stage 4
Stage 5
Dendrites
Spines
Establishment of polarity
(mid to late embryonic cerebral cortex)
Basal
Axon
Leading process
(future dendrite)
Basal
CP
Bipolar cell
Dendrites
Retinal
ganglion
cell
Neuroepithelial
cell
Apical
IZ-U
Multipolar cell
IZ-L
SVZ
VZ
Intermediate (basal)
progenitor cell
Trailing process
(future axon)
Radial glial cell
Apical
D
996
Locomotion
(radial glial cell-dependent)
Somal translocation
(radial glial cell-independent)
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Minor neurites
B
them. One day after seeding, neurons begin to show polarized morphology: several minor neurites and one neurite
that is much longer than the others (stage 3). The longest
neurite eventually becomes the axon. Therefore, the earliest
event of neuronal polarization in cultured neurons is axon
initiation. Then, within 7 days, the remaining neurites develop into dendrites (stage 4). Finally, the neurons form
NAMBA ET AL.
premature dendritic spines and axonal branches (stage 5).
Notably, in the absence of additional extracellular signals,
neurons determine their polarity in a stochastic manner.
B. Neuronal Polarization In Vivo
In mid to late cerebral cortical development, most of the
pyramidal neurons are generated from neural stem cells
defined as radial glial cells in the ventricular zone (VZ)
(213, 248, 333) via intermediate (basal) progenitor cells in
the subventricular zone (SVZ) (214, 249, 369). Because the
intermediate progenitor cells lose their polarity, the neurons
cannot inherit their polarity and thus must “establish” their
polarity again to become mature neurons. First, the newly
generated neurons extend multiple neurites that are identical to those in stage 2 cultured neurons, at which point they
are defined as MP cells in the lower portion of the intermediate zone (IZ) (119, 214, 237, 249, 287, 329). Almost
60% of MP cells initially extend a tangential trailing process, which ultimately develops into an axon, and then generate a leading process, which later develops into a dendrite
(119, 237). These neurons are defined as bipolar (BP) cells.
Fewer than 30% of MP cells initially generate a leading
process and then extend a trailing process. BP cells migrate
toward the cortical plate (CP) and develop into mature
pyramidal neurons. The MP-to-BP transition is a critical
step in neuronal polarization in vivo (119, 158, 233, 237).
In addition to axon-dendrite formation, neuronal polarization in vivo includes a different, but not completely separate, process: migration (231, 283) (FIGURE 1). Generally,
leading process formation is essential for migration toward
the proper destination. Therefore, the MP-to-BP transition
is also critical for neuronal migration (158, 167). Neuronal
migration in vivo is roughly separated into two modes:
somal translocation and locomotion (230). During the
somal translocation mode, neurons extend long processes
toward the direction of movement, and they then translocate their somata using these processes. After the translocation of the somata, the remaining process at the opposite
end will retract. In contrast, migrating neurons during the
locomotion mode extend relatively short leading processes.
Migrating neurons extend the leading process a short distance toward the direction of movement and then translocate the somata. The neurons repeat this cycle during locomotion. In the early developing cerebral cortex, most neurons migrate according to the somal translocation mode.
During the mid to late embryonic stages, neurons migrate
from the IZ to the CP according to a locomotion mode and
then convert to the somal translocation mode at the end of
migration. In general, somal translocation is radial gliaindependent, whereas locomotion is radial glia-dependent
(230).
C. Microenvironment of Neuronal
Polarization
Multiple cellular processes, such as cell migration and proliferation, occur within specific microenvironments in vivo.
The development of cells in vivo is distinct from in vitro cell
FIG. 1. Polarization process of neurons in cultured conditions and neurons in vivo. A: stage 1 neurons form several thin filopodia. Stage 2
neurons form multiple minor processes. Stage 3 neurons possess one long neurite (immature axon) and several minor processes. The longest
neurite ultimately develops into a mature axon. Therefore, the initial event of neuronal polarization in cultured neurons is axon initiation. Then,
within 7 days, the remaining neurites develop into dendrites (stage 4). Finally, the neurons form premature dendritic spines and axonal branches
(stage 5). B: retinal ganglion cells are directly derived from the retinal neuroepithelial cells. The apical process of the daughter cell eventually
develops into a dendrite, and the basal process that contacts the basal lamina becomes an axon. C: in the mid to late embryonic cerebral cortex,
most pyramidal neurons are generated from the radial glial cells in the ventricular zone (VZ) through intermediate (basal) progenitor cells in the
subventricular zone (SVZ). Then, the newly generated neurons in the lower part of the intermediate zone (IZ-L) extend multiple neurites. Most
of the multipolar cells first extend the trailing process tangentially and then generate the leading process. Finally, the multipolar cells transform
into bipolar cells and migrate toward the cortical plate (CP) while in contact with radial glial cells via the upper part of the intermediate zone (IZ-U).
In most cases, the initial step of neuronal polarization is the formation of the trailing process (future axon), as in cultured neurons.
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In contrast to cultured neurons, there are several types of
polarization processes in each neuronal subtype, brain region, and developmental stage in vivo. Polarization processes in vivo are roughly separated into the “inheritance of
polarity” mode and the “establishment of polarity” mode
(FIGURE 1). One of the best examples of the “inheritance of
polarity” mode is vertebrate retinal ganglion cells (and possibly neurons in the early developing cerebral cortex). Because
these neurons are directly generated from neuroepithelial cells,
they can inherit the apico-basal polarity of the neuroepithelial
cells (275). In retinal ganglion cells, the dendrites and axon are
differentiated from the apical process and the basal process,
respectively (383). The multipolar (MP) cells do not appear
during this developmental process. Interestingly, impairment
of the retinal neuroepithelial cell polarity induces the abnormal emergence of MP cells (383).
Most in vivo cells undergo the same initial step of neuronal
polarization as cultured neurons: trailing process (future
axon) formation (119, 237). Recently, several researchers
discovered a new type of basal (intermediate) progenitor
cell termed basal radial glial cells, specifically in animals
with gyrified brains (33, 76, 198, 338). This type of progenitor cell possesses clear polarity even when the cell body is
detached from the apical surface. Because these cells are
thought to be involved in brain evolution, it may be interesting
to analyze the polarization process in neurons derived from
the basal radial glia cells.
NEURONAL POLARITY SIGNALING
Upper IZ
(TAG-1 negative)
neuronal polarization in vivo (237). As described in the
previous section, the newly generated neurons develop MP
morphology in the lower IZ. In the lower IZ, there are many
tangentially oriented axons, called pioneering axons, arising from the early-born cortical neurons. Once one of the
neurites of a MP cell “touches” a pioneering axon, this
interaction induces the stabilization and rapid extension
(“go”) of the contacting neurite. The contacting neurite
eventually becomes an axon. TAG-1 (transient axonal glycoprotein-1), a member of the immunoglobulin superfamily
of cell adhesion molecules, is involved in this process (237).
One possible mechanism underlying this process is that
physical contacts increase the tension of immature neurites.
Mechanical tension induces neurite elongation, thereby
specifying the axon (187). MP cells repeatedly extend and
retract immature neurites, which could stabilize a single
neurite via cell-to-cell interactions, thereby increasing neurite tension and inducing axon specification. Another possible mechanism underlying this process is that the cell-cell
interactions activate signaling molecules that promote axon
specification. The downstream signal of TAG-1 is discussed
in the next section. Both of these mechanisms are likely
involved in axon specification according to the “Touch and
Go” mechanism.
The importance of the microenvironment for neuronal polarization in vivo raises a question: How do neurons in
culture establish their polarity without that microenvironment? The answer is very simple: the neurons create certain
microenvironmental cues by themselves. Neurons in culture
secrete neurotrophins such as brain-derived neurotrophic
factor (BDNF) and neurotrophin (NT)-3 (106, 233). Neurotrophins act as autocrine or paracrine microenvironmental cues for neuronal polarization in culture (233). Given
that neurons determine their polarity in a stochastic manner, these auto- or paracrine factors supply basic stimuli to
each minor process. Without neurotrophin signals, neurons
fail to polarize (233). This result suggests that microenviromental cues are essential for neuronal polarization even in
culture. The microenvironmental signals in vivo are more
Radial glia-neuron
interaction via N-cadherin
Auto- or paracrine action
of neurotrophins
Migration
Pioneering axon
Leading process
formation
Axon
specification
Lower IZ
(TAG-1 positive)
998
Neuron-neuron
interaction via TAG-1
Gradient of TGF-β
Basal process
of radial glial cell
FIG. 2. The microenvironment for neuronal polarization in vivo. Neurotrophins such
as BDNF and NT-3 act in an autocrine or
paracrine manner to regulate neurite formation (blue). A TGF-␤ concentration gradient along the radial axis regulates the radial
axon extension (green). The interaction between multipolar cells and pioneering axons
from early-born neurons is critical for neuronal polarization (magenta). Once one of
the neurites of a multipolar cell contacts
the pioneering axons through TAG-1, the
neurite extends rapidly and eventually becomes an immature axon. The interaction
between radial glial cells and neurons regulates the formation of the leading process
and neuronal migration (orange).
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culture conditions and is precisely regulated by environmental cues such as secreted factors, the extracellular matrix (ECM), and neighboring cells (83, 143). Immature neurons develop within a heterogeneous environment that also
contains secreted factors, the ECM, radial glial cells, and
other neurons (277). Recent studies have demonstrated the
involvement of several microenvironmental factors in neuronal polarity in vivo (FIGURE 2). Secreted factors including
neurotrophins (233) and transforming growth factor ␤
(TGF-␤) (376) regulate neuronal polarization in vivo in an
autocrine or paracrine manner (see below). ECM molecules
also regulate neuronal polarization. In the developing retina, retinal ganglion cells are generated directly from neuroepithelial cells. Shortly after cell division, the retinal ganglion cells reextend a basal process toward the basal surface, where laminin is expressed. Once the basal process
contacts laminin, the process is stabilized and ultimately
develops into an axon (276). Another microenvironmental
cue is cell-to-cell interaction (158, 168, 237). One such
example is the radial glial scaffold, which is involved in
neuronal migration. As described above, BP cells in the
upper portion of the IZ migrate toward the CP according to
the locomotion mode. Several types of junctions, such as
gap junctions and N-cadherin-mediated adherens junctions, are thought to be involved in this process (81, 158,
168). Interestingly, recent studies suggest that N-cadherinmediated cell adhesion is important not only for neuronal
migration but also for the MP-to-BP transition (92, 158,
168). Inhibiting the N-cadherin-mediated radial glial cellneuron interaction impairs the MP-to-BP transition. In addition, leading process formation is abnormal in neurons
transfected with a dominant-negative mutant N-cadherin
(92). These results indicate that N-cadherin-mediated interaction may regulate the MP-to-BP transition, particularly
leading process formation. Although these studies suggest
that the radial glial cell-neuron interaction is also important
for neuronal polarization (92, 158), this connection remains to be proven. Another example of a cell-to-cell interaction is the interaction between pioneering axons and neurons. Recently, we proposed the “Touch and Go” model for
NAMBA ET AL.
complicated than in culture. These microenvironments are
differentially expressed in space and time and might be
important for the location and timing of neuronal polarization (axon initiation).
III. MECHANISMS OF NEURONAL
POLARIZATION
Although many researchers are now interested in the mechanisms of neuronal polarization in vivo, much of the basic
knowledge comes from studies using dissociated neurons.
Therefore, we begin by reviewing some fundamental mechanisms that are presumably shared across in vivo and in
vitro systems and then discuss the current knowledge about
signaling pathways for neuronal polarization in vivo.
1. Cytoskeletal organization
One principal element of neuronal polarity is cytoskeletal
organization. Almost all signals modulate cytoskeletal organization, thus determining cell morphology and motility
(11, 19, 193, 367). In particular, growth cone dynamics are
regulated by two cytoskeletal components, microtubules
and actin filaments (59, 196) (FIGURE 3). The growth cone is
Dendrite
Mixed microtubule orientation
Axon
Plus-end-distal microtubule orientation
Distal
Distal
+ – + –
+ + + +
– + – +
– – – –
Proximal
Proximal
A) MICROTUBULES. Microtubules are important regulators of
the polarity of many cell types (73, 193, 323). Microtubules are composed of ␣-tubulin and ␤-tubulin heterodimers and consist of two distinct ends: the plus and
minus ends (FIGURE 4). The plus end is a crucial site for
tubulin polymerization. The minus end is often anchored
to a microtubule-organizing center (MTOC), such as the
centrosome and the Golgi complex.
The first two reports that described the orientation of microtubules in neurons were published in 1981 (39, 123).
These studies independently reported that the microtubules
in axons are uniformly oriented, with distally directed plus
Growth cone
Microtubule
C-domain
T-zone
Dendrite
Axon
P-domain
Actin
FIG. 3. The cytoskeletal structures in neurons. The dendrites and axons have different microtubule orientations; dendritic microtubules show mixed orientation, and axonal microtubules show plus-end-distal orientation.
Growth cones are divided into three regions based on their cytoskeletal components. The central (C) domain
mainly contains microtubules, and the peripheral (P) domain is enriched in actin filaments and contains filopodia
and lamellipodia-like structures. The transition (T) zone contains an acto-myosin contractile structure.
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A. Basic Knowledge About Neuronal
Polarization: Lessons From Dissociated
Neurons
divided into three regions based on their cytoskeletal components (196). The central (C) domain mainly contains microtubules, and the peripheral (P) domain is rich in actin
filaments and contains filopodia and lamellipodia-like
structures. Between these domains, there is an acto-myosin
contractile structure termed the transition (T) zone. The
three steps of growth cone dynamics that result in neurite
outgrowth are as follows: protrusion, engorgement, and
consolidation (99). Membrane protrusion is primarily
driven by filamentous actin (F-actin) polymerization. Engorgement results from microtubule polymerization and
microtubule-mediated transport of membranous organelles
and vesicles. Consolidation is the process of F-actin depolymerization at the neck (proximal region). In this section,
we focus on microtubule and actin filament dynamics.
NEURONAL POLARITY SIGNALING
A
MTOCs (centrosome, Golgi complex, etc.)
Minus-end binding proteins
(ninein, CAMSAPs, etc.)
MAPs (MAP2, tau, etc.)
Minus end
Plus end
+TIPs (EBs, CLASP, etc.)
α
β
α
β
GTP-tubulin
GDP-tubulin
B
Luminal surface
Am
Am
α
Tyrosinaltion (TTL, etc.)
Detyrosination (possibly CCP2, etc.)
Ac
β
Am
Am
D
E
Y
E
Δ2-tubulin
Δ3-tubulin
E
G
E
E
E
E
E
Q
G
E
E
G
Outer surface
E
E
E
A
G
E
E
E
G
E
E
F
G
E
E
Polyamination
(TG1, 2, 3, 6, etc.)
G
E
G
G
G
Polyglutamylation (TTLL1, 4, 5, 6, 7, 11, 13, etc.)
Deglutamylation (CCP1, 4, 5, 6, etc.)
Polyglycylation (TTLL 3, 8, 10, etc.)
Deglycylation (unkown)
C
Actin-severing proteins (ADF/cofilin, etc.)
Branched actin nucleators (Arp2/3, etc.)
Pointed end
Barbed end
Barbed-end protector proteins
(Ena/VASP, etc.)
ADP-actin
ATP-actin
ends in the peripheral nervous system, such as the frog
olfactory nerve and the cat lumbar colonic nerve. Then,
Baas and colleagues (16, 17) found that the orientation of
microtubules in axons is plus-end-distal and that the orientation of microtubules in dendrites is mixed in cultured rat
hippocampal neurons. Based on their results and those of
modern studies using EB3-GFP to label the plus ends in
living cells (319, 368), microtubule orientation during neuronal polarization was analyzed in detail. At stage 2, microtubule orientation in minor neurites is uniformly plus-enddistal. At stage 3, microtubule orientation in axons is plusend-distal and remains plus-end-distal in minor neurites,
identical to that in stage 2 neurons. As dendrites develop
1000
(after stage 4), both plus- and minus-end-distal microtubules are detected in dendrites; however, only plus-enddistal microtubules are detected in axons (FIGURE 3). Alternatively, microtubule orientation during neuronal polarization in vivo is complex (286). A recent study using the
EB3-GFP labeling method and an embryonic cortical slice
culture system demonstrated that the orientation of microtubules in MP cell neurites is plus-end-distal, identical to
that of stage 2 cultured neurons, whereas the orientation of
microtubules in the trailing process (nascent axon) is mixed
(287). Because the orientation of microtubules in mature
axons in the cerebral cortex remains unclear, we cannot
determine whether this difference is dependent on the envi-
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Acetylation (αTAT, etc.)
Deacetylation (HDAC6, etc.) Ac
FIG. 4. Structure and dynamics of microtubules and actin filaments. A: microtubules are composed of ␣-tubulin and ␤-tubulin heterodimers and have dynamic plus
ends and less dynamic minus ends. The
⫹TIPs bind to the plus ends. The minus end
is anchored to a microtubule-organizing
center (MTOC) or associates with minusend-binding proteins. The lattice of microtubules is associated with microtubule-associated proteins (MAPs). B: tubulin modifications and proteins responsible for each
modification. ␣TAT, ␣-tubulin N-acetyltransferase; CCP, cytosolic carboxypeptidase;
HDAC, histone deacetylase; TG, transglutaminase; TTL, tubulin tyrosine ligase;
TTLL, TTL-like. C: actin filaments are composed of actin and have barbed ends and
pointed ends. Actin-severing proteins enhance actin disassembly. The branching of
actin filaments is catalyzed by the actin nucleator Arp2/3. The barbed end interacts
with barbed-end-protecting proteins.
NAMBA ET AL.
ronment, such as cultured conditions versus in vivo conditions or peripheral versus central, or is simply dependent on
axon maturation.
Previous studies demonstrated that the stability of microtubules in axons is different from that in minor neurites or
dendrites (20, 116, 368). The posttranslational modification of tubulin, primarily ␣-tubulin, affects microtubule dynamics (90). Several types of tubulin posttranslational modifications have been identified: tyrosination, acetylation,
Tubulin acetylation is a result, not a cause, of microtubule stabilization (363). Unlike most posttranslational
modifications of tubulin, which occur outside the microtubule, acetylation occurs on residues within tubulin
(90). ␣-Tubulin is known to be acetylated by MEC17/
␣TAT in C. elegans neurons (3, 306) and also in the
mouse central nervous system (161) and to be deacetylated by histone deacetylase 6 (HDAC6) and by the
NAD-dependent deacetylase sirtuin-2 (146, 250). Although increasing tubulin acetylation via trichostatin A
treatment or HDAC6 knockdown results in decreased
axon elongation, HDAC6 overexpression does not affect
axon formation (336). Interestingly, trichostatin A treatment or HDAC6 knockdown impairs axon initial segment (AIS) formation; thus tubulin acetylation may be
involved in maintaining neuronal polarity.
Recently, a study suggested a novel mechanism for modulating microtubule stability in axons (316). This study
showed that polyamination of tubulin, which is mediated
by transglutaminase, increases its stability. Because Ca2⫹
activates transglutaminase, neurotrophin-induced Ca2⫹ release might regulate axonal microtubule stability via polyamination. Although transglutaminase regulates neurite extensions in neuroblastoma cells, its functions in neurons
and in vivo remain unknown.
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The orientation of microtubules primarily relies on the
MTOCs (27, 197). One major MTOC is the centrosome.
Typically, centrosomes are composed of two centrioles and
pericentriolar material, such as a ␥-tubulin ring complex
(␥-TuRC) (197). ␥-Tubulin nucleates microtubule polymerization; therefore, microtubules are tethered to the
␥-TuRC via their minus ends. In addition, the aPKC-dependent activation of Aurora-A kinase phosphorylates Ndel1,
most likely inducing nucleation at MTOCs (223). This sequential event induces neurite elongation in dorsal root
ganglia neurons. Although the position of the centrosome
has been proposed to be a primal regulator of axon specification (56, 190), recent studies using cultured neurons and
in vivo models demonstrated that the position of the centrosome is a result, not a cause, of axon specification (287,
320). These studies showed that axon specification occurs
independently from the position of the centrosome. In addition, axon elongation occurs normally in cultured neurons with experimentally ablated centrosomes (320). The
finding that noncentrosomal microtubules are dominant in
neurons suggests the pivotal function of noncentrosomal
MTOCs and microtubules (16). In general, the noncentrosomal microtubule array is generated via three mechanisms:
release from centrosomes, capture by noncentrosomal
MTOCs, or interaction with minus end-binding proteins
followed by transport and assembly (27). Microtubules are
released from centrosomes via the microtubule-severing
protein katanin (118, 208). Inhibiting katanin function in
cultured neurons results in the impairment of axon extension (2). Subsequently, the noncentrosomal microtubules
are captured by noncentrosomal MTOCs, such as the cortical membrane, the nuclear envelope, and the Golgi complex. In neurons, it has been shown that the Golgi complex
in dendrites, referred to as the Golgi outpost, acts as an
MTOC and regulates dendritic branching (255). However,
Golgi outposts and other MTOCs, such as the ␥-TuRC,
have not been found in axons (18). Thus the free minus ends
might be capped by minus end-binding proteins, such as
ninein (23, 34, 216), or by the calmodulin-regulated spectrin-associated protein (CAMSAP)/Nezha/Patronin family
of proteins (22, 155, 334, 375) to stabilize the microtubules. Finally, the noncentrosomal microtubules are transported to the proper subcellular region and aligned in their
proper orientation. However, the mechanism by which microtubule orientation is determined remains unclear.
polyglutamination, polyglycylation, palmitoylation, phosphorylation, and polyamination (FIGURE 4). Of these modifications, the roles of tyrosination, acetylation, and polyamination of tubulin have been analyzed in detail in neurons; thus, in this review, we focus on these modifications.
The COOH terminus of ␣-tubulin is tyrosinated by tubulin
tyrosine ligase (TTL) (229, 279, 296) and presumably
detyrosinated by CCP2 (AGBL2) (284). The tyrosination of
tubulin recruits microtubule depolymerizers, such as kinesin-13 (Kif2A, 2B, and 2C), thereby contributing to the
shorter lifetime of tyrosinated microtubules (266). TTL
knockout mice exhibit abnormal layer formation and axon
formation in the cerebral cortex (71), and enhanced axon
specification and elongation are detected in TTL knockout
cultured neurons (178). In contrast, TTL overexpression in
cultured neurons inhibits axon outgrowth (271). In addition, Kif2A knockout mice also exhibit abnormal layer formation in the cerebral cortex, and enhanced axon elongation and branching are detected in Kif2A knockout cultured
neurons (136). Therefore, the tubulin tyrosination/detyrosination cycle is an important regulator of neuronal polarization, including migration and axon specification. Interestingly, removing the penultimate glutamate residue from
␣-tubulin forms ⌬2-tubulin, which cannot be tyrosinated,
resulting in exit from the tyrosination/detyrosination cycle
(261). Microtubules composed of ⌬2-tubulin are the major
component of the brain and neurons (262). Unlike detyrosinated tubulin, ⌬2-tubulin is abundant in growth cones.
NEURONAL POLARITY SIGNALING
B) ACTIN FILAMENTS.
The dynamic organization of the actin
cytoskeleton directs the morphology of growth cones (59,
196). Actin filaments contain two distinct ends: the barbed
and pointed ends (FIGURE 4). In the growth cone, ATP-actin
is added to the barbed end, which is directed toward the cell
membrane, and ATP hydrolysis and ADP-actin disassembly
occur at the pointed end. Therefore, these actin filament
dynamics cause retrograde actin flow from the peripheral
region to the central region of the growth cone. The formation of highly branched arrays of actin filaments is important for lamellipodia organization (326). The branching of
actin filaments is catalyzed by the actin-nucleating Arp2/3
complex (5). The Arp2/3 complex binds to the side of the
actin filament and initiates new actin filament assembly.
Although much is known regarding the roles of actin filament dynamics in the migration of nonneuronal cells (193),
our current understanding of the mechanism by which actin
filament dynamics regulate neuronal polarity is far from
complete. In this review, we focus on the role of actin filaments in axon formation.
A previous study demonstrated that the actin cytoskeleton
is more dynamic in the growth cone of the future axon than
in other minor neurites (37). Cultured neurons treated with
cytochalasin D, an actin-depolymerizing drug, extend multiple axons. These results suggest that an appropriate level
of actin instability at growth cones is critical for axon formation. Consistent with this insight, the actin-severing protein family ADF/cofilin (25, 242) regulates axon formation
1002
by creating space for microtubules to penetrate into the
peripheral regions of growth cones (77). In contrast, it has
been previously shown that the Ena/VASP family of barbed
end protector proteins (96, 112) is required for neurite formation by regulating actin dynamics and by promoting actin bundle formation in the peripheral regions of growth
cones of neurons cultured on poly-D-lysine (PDL) (111).
Actin bundle formation is important for microtubule penetration into the peripheral regions of growth cones. In Ena/
VASP mutant mice, axon formation in the cerebral cortex is
severely defective (184). Activating integrin signals rescues
the phenotypes of Ena/VASP mutant neurons. When neurons are cultured on laminin, these neurons require Arp2/3
rather than Ena/VASP for axon elongation (111). Inhibiting
Arp2/3 in neurons increases axon elongation in the presence
of PDL (267) but inhibits axon elongation in the presence of
laminin (111). Therefore, the coordinated regulation of actin filament and microtubule dynamics is essential for axon
formation; however, neurons may select the machinery for
this process based on the surrounding microenvironment.
2. Intracellular transport
The intracellular transport of cytosolic proteins, membranous structures, and organelles is a critical regulator of cell
polarity. Motor proteins, including the myosin, dynein, and
kinesin families of proteins, are responsible for the anterograde and retrograde transport of cargo (131, 238, 295). In
general, myosins are actin-dependent motors, whereas dyneins and kinesins are microtubule-dependent motors. In
this section, we survey the mechanism by which cargo is
selectively transported to the appropriate subcellular region
by motor proteins (FIGURE 5).
A) MOTOR PROTEINS. Fifteen kinesin protein families, consisting of 45 mammalian kinesin superfamily proteins (KIFs),
exist. The most characterized kinesin is kinesin-1. Kinesin-1
is a complex of two kinesin heavy chains (KHCs, also called
KIF5), and two kinesin light chains (KLCs) (32, 183). KIF5
contains a motor domain and an alpha-helical coiled-coil
domain. Of the 45 identified KIFs, KIF1, KIF2, KIF3, KIF4,
KIF5, KIF13, KIF17, KIF26, KIF20B, and KIFC2 are
known to play important roles in cargo transport in neurons. Kinesins move toward the plus-end of microtubules,
and microtubules are plus-end-distal in axons; thus kinesins
are responsible for most of the anterograde cargo transport
in axons (131, 238, 295).
The kinesin family of proteins selectively transports various
types of cargos, such as organelles and cytosolic, membrane-bound, and cytoskeletal proteins. For example, the
anterograde transport of mitochondria into axons is mediated by kinesin-1 (97, 110, 322). Vesicles containing the
neurotrophin receptor TrkB are transported toward the distal end of the axon by kinesin-1 (12). Kinesin-4 (KIF4)
transports membrane proteins such as the cell adhesion
molecule L1, which is involved in various developmental
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Another class of regulators of microtubule stability is microtubule-associated proteins (MAPs). In neurons, MAP2,
a dendrite-specific MAP, and tau, an axon-specific MAP,
prevent the interaction between microtubule-severing proteins and microtubules, thereby contributing to microtubule stability (68, 160). These proteins may also contribute
to axon-dendrite polarity in cultured neurons (42, 43, 74).
A more recent study suggests that one polarity protein,
partitioning-defective 3 (Par3), regulates the stability of microtubules (47). Briefly, Par3 oligomerization induces conformational changes in Par3; subsequently, Par3 binds to
and stabilizes microtubules. The details of this function of
Par3 are described below. Doublecortin (Dcx) has been
identified as a causal gene of human X-linked lissencephaly
and double cortex syndrome (60, 98). Dcx is expressed in
migrating neurons (80) and regulates migration and the
MP-to-BP transition in the developing cerebral cortex (21,
203, 292). Dcx binds to the lattice of microtubules and
promotes microtubule nucleation and stability (79, 219).
The interaction between Dcx and microtubules is mediated
by “Dcx domains,” which may cross-link two tubulin subunits on the same or different microtubules. Dcx activity is
partially regulated by Dcx phosphorylation. Dcx phosphorylation by several kinases, such as Cdk5, cAMP-dependent
protein kinase (PKA), and MARK, reduces the affinity of
Dcx to microtubules (104, 293).
NAMBA ET AL.
CaMKII
ERK1/2
NR2B
Par6
P
Mint1 (Lin10)
aPKC
KIF17
KIF3
GAP
Rab3
GDP
DENN/
MADD
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GAP
Rab3
GTP
P
Par3
Rab27
GTP
Rab27
GDP
TrkB
Slp1
CRMP-2
KIF1A
KIF1Bβ
KIF5
Mitochondria
Mitochondria
Miro
Miro
Milton
+Ca2+
KIF5
Milton
+Ca2+
KIF5
FIG. 5. Intracellular transport in neurons. Motor proteins, cargos, and cargo adaptors and their loading and
unloading mechanisms. P indicates phosphorylation.
processes in neurons, toward the distal end of the axon
(265, 289). Cytoskeletal proteins, such as neurofilaments
and tubulin, are transported into axons by kinesin-1 (174,
177, 340, 370, 371). Selective transport by kinesins is important for synaptic transmission in mature neurons. Kinesin-3 regulates the transport of synaptic vesicle precursors
into axon terminals (114, 246, 253, 382).
The kinesin family of proteins also transports specific proteins that regulate axon formation. For instance, Par3 is
transported to the tips of developing axons by kinesin-2
(KIF3A) (88, 244). Collapsin response mediator protein-2
(CRMP-2) is transported to the distal ends of nascent axons
by kinesin-1 through binding with KLC (12, 166, 174,
348). Both Par3 and CRMP-2 are predominantly localized
at the distal ends of axons and play pivotal roles in axon
specification. Disrupted-in-schizophrenia 1 (DISC1), a susceptibility gene for major psychiatric disorders, including
schizophrenia (50, 153), is transported by kinesin-1 to axon
growth cones (309, 339). This transport is important for
axon elongation. Kinesin-6 (KIF20B), a mitotic kinesin, interacts with Shootin1 and regulates its transport toward the
tips of axons (290). In cooperation with L1 and actin, Shootin1 induces axon specification in cultured neurons and in
vivo, generating a driving force in growth cones (307, 343,
345). This traction force is regulated by the PAK-mediated
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1003
NEURONAL POLARITY SIGNALING
phosphorylation of Shootin1 (343). Because Shootin1 is
transported toward the tips of axons by a growth cone-like
structure, termed wave, the mechanism by which neurons
select between these two independent mechanisms to transport Shootin1 is interesting. PIP3-containing vesicles are
transported by guanylate kinase-associated kinesin
(GAKIN)/KIF13B (139). GAKIN/KIF13B is known to regulate axon specification in cultured neurons (139), and its
activity is regulated by MARK2 (379).
The dynein family members are classified into two groups:
axonemal and cytoplasmic dyneins (353). Cytoplasmic dyneins move along the microtubules toward the minus-end;
thus these motors are referred to as minus-end-directed motors. Cytoplasmic dyneins contain two dynein heavy chains,
light intermediate chains, and a complex of intermediate
and light chains. Because detailed information regarding
the structure and basic function of dyneins is reviewed in an
excellent article (353); in this review, we focus on the role of
dyneins in neuronal development. The activity of dyneins is
known to be regulated by dynactin and LIS1/NDEL (297,
312, 360). LIS1 was originally identified as a causal gene of
type I lissencephaly (282). The suppression of dynein or
LIS1 expression impairs neuronal migration in the developing cerebral cortex (133, 335, 346). On the bases of these
results, a dynein-based neuronal migration model has been
proposed (347). Briefly, dynein-based neuronal migration
consists of three steps. First, leading process extension occurs. As the leading process elongates, the centrosome
moves toward the leading process and causes “swelling” at
the proximal region of the leading process. Second, the
nucleus translocates to this swelled region (defined as
nucleokinesis). Finally, the cell soma catches up to the nucleus. Because it had been believed that the centrosome is
always ahead of the nucleus and that the nucleus is surrounded by centrosome-derived microtubules, the forward
movement of the nucleus was thought to be induced by the
dynein complexes. However, recent studies demonstrated
that the centrosome-nucleus positioning is sometimes reversed (287, 352). Therefore, the simplified dynein-based
model requires revision (286). One possible mechanism is
based on the acto-myosin-dependent driving force to translocate the nucleus (29, 294).
B) THE MECHANISMS OF SELECTIVE TRANSPORT.
Selective targeting of different cargo to a specific subcellular region is
1004
C) PREFERENCES OF KINESINS.
Although most kinesins move
toward the plus-end of microtubules, dendrite- or axonspecific kinesins have been identified. In polarized neurons
(stage 3 and stage 4 neurons), the motor domains of KIF5,
KIF3, KIF13A, and KIF21A preferentially accumulate at
the tips of axons rather than in dendrites (144, 157, 234).
Although KIF17 immunoreactivity is selectively localized to
dendrites in vivo and in cultured neurons (300), the motor
domain of KIF17 accumulates in axons (144). Because the
tail domains of KIF proteins regulate the motor activity of
KIFs by interacting with the head domain (63), the tail
domain likely regulates the selective localization of KIF proteins in some cases. In addition, KIF5 transports the
AMPA-R subunit GluR2 to dendrites by interacting with
the adaptor protein glutamate receptor-interacting protein
(GRIP1) (301), suggesting that the interaction between the
cargo and the motor protein plays an important role in
selective transport (132). In nonpolarized neurons (stage 2
neurons), the motor domains of KIF5 and KIF17 accumulate in a subset of neurites (144). However, the motor domains of other KIF proteins fail to accumulate or localize to
most neurites (144). Therefore, how do KIF proteins determine their selectivity? The evidence suggests that tubulin
posttranslational modifications determine the axon-dendrite preference of KIFs. The KIF5 motor domain has been
suggested to preferentially recognize axonal microtubules
(178). Interestingly, the affinity of KIF5 for microtubules is
enhanced or reduced by tubulin posttranscriptional modifications, such as acetylation and tyrosination, respectively
(178, 280). As described above, acetylated and detyrosinated microtubules are enriched in axons, whereas tyrosinated microtubules are abundant in dendrites of polarized
neurons (116, 368). Therefore, tubulin acetylation at axons
may induce the selective transport of KIF5 to axons, and
tubulin tyrosination may exclude KIF5 from dendrites
(178, 280). Previous studies demonstrated that taxol treatment, which induces the acetylation, detyrosination, and
polyglutamination of tubulin, results in the accumulation of
the KIF5 motor domain at the tips of minor neurites as well
as at the tips of axons (116). However, specifically increasing tubulin acetylation using trichostatin A or tubacin does
not alter the axon-dendrite preference of KIF5 (116). A
more recent study suggests that the KIF5 motor domain
preferentially binds to GTP-tubulin (235). These results indicate that the axon-dendrite preference of KIF5 is controlled by multiple posttranslational modifications and/or
by binding to GTP/GDP-tubulin.
D) CARGO ADAPTORS. Most kinesins interact with their specific
cargos via specific cargo adaptors (131, 238, 295) (FIGURE
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The kinesin family of proteins also regulates cargo transport
into dendrites. In dendrites, there are several types of neurotransmitter receptors. Of these, the N-methyl-D-aspartate
receptor (NMDA-R), ␣-amino-3-hydroxyl-5-methyl-4-isoxazole-propionate receptor (AMPA-R), and ␥-aminobutyric
acid receptor (GABAR) are transported by kinesin-1 (KIF5)
or kinesin-2 (KIF17) (300, 301, 350). In addition, postsynaptic proteins, such as postsynaptic density-95 (PSD-95),
are transported by kinesin-3 (KIF1B) (217).
crucial for the polarized distribution of molecules. Selective transport is established via three mechanisms: the
preferential and selective binding of motor proteins to
cargo adaptors, the loading and unloading of cargo, and
selection at the AIS (see below) (131, 238, 295).
NAMBA ET AL.
5). Several cargo adaptors involved in selective transport
have been identified. In this review, we discuss several wellcharacterized cargo adaptors, c-Jun N-terminal kinase
(JNK)-interacting proteins (JIPs), nuclear distribution gene
C homolog (NUDC), Par3, DISC1, and CRMP-2.
NUDC is a subunit of the dynein complex and is known to
regulate nuclear movement during interphase in Aspergillus
nidulans (256). The NUD family of proteins, including
NUDC, NDEL1, and LIS1 (NUDF), regulates nuclear
translocation during neuronal migration by interacting
with dynein (14, 44, 225, 310). NUDC binds to KLC,
thereby indirectly linking the dynein complex to KIF5 and
regulating the anterograde transport of dynein in fibroblasts (372). However, the roles of NUDC in kinesin-dependent cargo transport in cultured neurons and also in vivo
remain unclear.
Par3 forms a complex with Par6 and aPKC and accumulates at the distal end of growing axons (244, 305). Because
Par3 interacts directly with kinesin-2 (KIF3A) (88, 244),
Par3 is a cargo adaptor between the Par complex and kinesin-2, thereby regulating the transport of the complex toward the tips of nascent axons.
DISC1 acts as a cargo adaptor for kinesin-1 (KIF5) by interacting with NDEL1, LIS1, growth factor receptor-bound
protein 2 (Grb2), Girdin, cAMP-specific 3=,5=-cyclic phosphodiesterase 4B (PDE4B), glycogen synthase kinase-3␤
(GSK-3␤), and fasciculation and elongation protein zeta-1
(Fez1) (70, 162, 172, 204, 212, 309, 339). DISC1 controls
the axonal transport of NDEL1/LIS1, Grb2, and Girdin
(70, 309, 339). These interactions are important for axon
elongation.
CRMP-2 was originally identified as a mediator of semaphorin-3A signaling in the nervous system (102) and is specifically localized to axons (148). CRMP-2 interacts with
the KLC of kinesin-1, thus regulating the axonal transport
E) LOADING AND UNLOADING OF CARGO. To selectively transport
specific cargos to specific regions, the critical events are the
loading and unloading of cargo (FIGURE 5). Previous studies
demonstrated that several signals, including protein kinases, Rab GTPases, and Ca2⫹, regulate the loading and
unloading of cargo (131, 238, 295).
The phosphorylation of cargo adaptors and kinesins has
been suggested to regulate the loading and unloading of
cargo from kinesins. GSK-3␤ phosphorylates CRMP-2 and
interrupts its interactions with both tubulin (378) and the
KLC of kinesin-1 (12), thereby inducing the unloading of
cargo from the cargo adaptor (CRMP-2) and kinesin-1.
GSK-3␤ activity might thus regulate CRMP-2 activity, indirectly controlling the anterograde transport of specific
cargos, such as tubulin, the Sra1-WAVE1 complex, and
Trks, during axon formation. GSK-3␤ also phosphorylates
KLC and may induce the degradation of KLC. Membranebound organelles are unloaded from kinesin-1 upon GSK3␤-dependent phosphorylation of KLC (222). Rho-associated kinase (Rho-kinase; ROCK, ROK), an effector of the
small GTPase RhoA, also phosphorylates CRMP-2,
thereby inhibiting the interaction of CRMP-2 with tubulin
and with the KLC of kinesin-1 (13).
Calcium/calmodulin-dependent protein kinase II (CaMKII)
phosphorylates the tail domain of KIF17 (kinesin-2) (107).
Kinesin-2 (KIF17) transports NMDA-Rs as cargo via the
adaptor proteins LIN10 or Mint1. The unloading of cargo
from KIF17 is regulated by KIF17 phosphorylation (107).
CaMKII also phosphorylates liprin-␣1, which is a cargo
adaptor for leukocyte antigen-related tyrosine phosphatase
(LAR-RPTP) (138). Phosphorylation of liprin-␣1 induces
its degradation via the ubiquitin-proteasome system,
thereby inducing the unloading of LAR-RPTP-containing
cargo from kinesin-3 (KIF1A). Because the activity of CaMKII
is regulated by NMDA-R-dependent Ca2⫹ influx, the unloading of the NMDA-R and other cargo from kinesins may
play an important role in the activity-dependent recruitment of NMDA-Rs and other molecules to the postsynaptic
membrane.
Activating the JNK signaling pathway leads to the dissociation of JIPs from KLC (kinesin-1), thereby inducing the
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JIPs are scaffolding proteins for the JNK signaling pathways
(61). JIPs also act as cargo adaptors to transport cargo into
axons (181). JIPs serve as bridges between KIF5 and cargos
such as phosphorylated ␤-amyloid precursor protein (APP)
(150) and apolipoprotein E receptor 2 (ApoER2), which is
a Reelin receptor (354). In Drosophila, JIP regulates axondirected cargo transport in a KIF5-dependent manner (35,
40, 140, 141). In contrast, JIP1 or JIP1/JIP2 knockout mice
do not exhibit any abnormalities in axon-directed cargo
transport (171, 366). Because there are four mammalian
isoforms of JIP, the differences between insects and mammals may be caused by the functional redundancy of the
mammalian JIPs. Knockdown of JIP1 inhibits polarization
of cultured neurons (54). In contrast, knockdown of JIP3
does not affect polarization but inhibits axon elongation in
cultured neurons and in vivo (145).
of tubulin heterodimers and the Sra1-WAVE1 complex as a
cargo adaptor (166, 174). Similarly, UNC-33, a homolog of
CRMP-2 in C. elegans, links tubulin heterodimers to KLC-2
of kinesin-1 (348). An additional study demonstrated that
CRMP-2 controls the transport of TrkB in a kinesin-1dependent manner (12). In this process, CRMP-2 links
TrkB to kinesin-1 through Slp1 and Rab27 (12). Therefore,
CRMP-2 acts as a cargo adaptor and thereby regulates axon
formation by transporting the critical proteins involved in
remodeling microtubules and actin filaments and in trafficking growth factor receptors.
NEURONAL POLARITY SIGNALING
unloading of cargo in Drosophila (141). The activation of
DLK and MKK7, which are components of JNK pathway
kinases, has been shown to inhibit the interaction between
kinesin-1 and APLIP1, a Drosophila homolog of JIP1. Although many studies have demonstrated that phosphorylation regulates the loading and unloading of cargo, as described above, only a few studies have demonstrated the
significance of the loading and unloading of cargo to neuronal polarity.
The Rab family of GTPases regulates vesicle trafficking at
various steps (87, 318, 331, 381). Rab GTPases serve as
molecular switches by cycling between their GDP-bound
inactive and GTP-bound active forms. Rab GTPase activity
is regulated by Rab GDP/GTP exchange factors (GEFs),
Rab GTPase-activating proteins (GAPs), and Rab guanine
nucleotide dissociation inhibitors (GDIs) (87, 318, 331,
381). Rab GEFs convert GDP-bound Rab GTPases to
their corresponding GTP-bound form, thus activating
Rab GTPases. In contrast, Rab GAPs convert the GTPbound form to the GDP-bound form, thus inactivating
Rab GTPases (87, 318). GTP-bound Rab GTPases interact with and activate their specific effectors. Previous
studies have shown that Rab GTPases regulate the loading and unloading of specific cargo from kinesins or from
cargo adaptors (10, 246).
A previous study suggested that Rab3 plays an important
role in the loading and unloading of synaptic vesicle precursors (246). GTP-bound Rab3, but not GDP-bound Rab3,
can interact with DENN/MADD. Because DENN/MADD
binds to kinesin-3 (KIF1A/KIF1B) as a cargo adaptor, only
GDP-bound Rab3 can link Rab3-containing vesicles to kinesin (246). Therefore, Rab3 activity regulates the loading
and unloading of cargo. Because a previous study suggested
that DENN/MADD acts as a GEF on Rab3 (373), the
DENN/MADD-Rab3 complex may sustain Rab3 activity
and facilitate the stable transport of Rab3-containing vesicles. Vesicle unloading may be regulated by Rab3-specific
GAPs. Interestingly, Rab3-GAPs regulate synaptic transmission (288), suggesting that Rab3 is important for vesicle
transport to regulate synaptic function.
1006
The motility and localization of mitochondria are regulated
by the intracellular Ca2⫹ concentration (45, 135). Mitochondria are transported by kinesin-1 (KIF5) via the
MIRO-Milton adaptor protein complex in Drosophila (97,
135) and in mammalian neurons (38, 82, 199, 313). Increased Ca2⫹ levels induce conformational changes in
MIRO through its EF hand motifs to bind to KIF5 (359).
This MIRO-KIF5 interaction interferes with the interaction
between KIF5 and microtubules, arresting mitochondrial
movement. Another potential mechanism of mitochondrial
unloading is that elevated Ca2⫹ concentrations may decrease the affinity of the MIRO-Milton complex for KIF5,
thereby inhibiting mitochondrial movement (200). Further
studies are required for understanding the precise mechanisms of the Ca2⫹-induced unloading of cargo.
3. Intracellular signal molecules
Various types of intracellular signaling molecules have been
identified as regulators of neuronal polarity. In this section,
we introduce the roles of the Rho family of small GTPases,
kinases, scaffolding proteins, transcription factors, and epigenetic modifiers in neuronal polarization (FIGURE 6).
A) THE RHO AND RAS FAMILIES OF SMALL GTPASES.
Rac1, Cdc42,
and RhoA, members of the well-understood Rho family of
small GTPases, play crucial roles in the morphogenesis of
various cells by reorganizing the cytoskeleton (113). Because Rho GTPases cycle between GDP-bound inactive and
GTP-bound active states, these molecules act as molecular
switches. The active forms of Rho GTPases interact with
and activate a variety of specific downstream effectors, and
they mainly regulate cytoskeletal reorganization.
Neurons electroporated with constitutively active (CA)
Rac1, dominant-negative (DN) Rac1, or Rac1 GEFs, such
as STEF/Tiam1 or P-Rex1, exhibit retarded migration and a
loss of leading and trailing processes, suggesting that the
balance of Rac activity is required for the MP-to-BP transition and migration in vivo (167, 179, 380). Because several
Rac isoforms are expressed in the neocortex, the defects in
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Recently, one study demonstrated that Par3 phosphorylation at serine 1116 by extracellular signal-regulated
kinase (ERK) disrupts the binding of Par3 to KIF3A (88).
Because phosphorylated Par3 accumulates at the tips of
axons in cultured neurons, this phosphorylation might
regulate the unloading of the Par complex from KIF3A.
The knockdown of Par3 impairs axon specification and
the MP-to-BP transition in cultured neurons and in vivo,
respectively. Unlike wild-type Par3, a phosphomimetic mutant of Par3 (S1116D) did not rescue the Par3 knockdown
phenotype. Therefore, the appropriate unloading of the Par
complex is critical for neuronal polarization both in cultured neurons and in vivo (88).
Rab27, the most similar isoform to Rab3, also regulates the
trafficking of a variety of secretory vesicles in many types of
cells, including neurons (86). Rab27 is localized to TrkBcontaining vesicles and regulates the anterograde transport
of Trks in axons (10). The GTP-bound form of Rab27 can
interact with an effector of Rab27, synaptotagmin-like protein 1 (Slp1). Slp1 directly binds to the cytoplasmic region
of TrkB in a manner dependent on activated Rab27. Slp1
also interacts with CRMP-2. Thus the Slp1/Rab27B/
CRMP-2 complex directly links TrkB-containing vesicles to
kinesin-1. Because the formation of this complex depends
on the GTP/GDP-binding state of Rab27, Rab27 may regulate the loading and unloading of TrkB-containing vesicles. Further studies are required to understand the mechanisms by which Rab27 activity is regulated in neurons.
NAMBA ET AL.
Neurotrophin
(BDNF, NT-3)
TrkR
Plasma membrane
Plexin
Neuron-neuron
interaction
TGF-β, BMP
Sema3A
NP-1
GPCR
TGF-βR
TAG1
Reelin
ApoER
VLDLR
Lyn
N-Cadherin
Dab1
p120
IP3
IP3
receptor
PLC
AC
sGC
Ca2+
cAMP
cGMP
DLK
CaMKK
PKA
PKG
MKK4/7
Ras
SMAD
Raf
PIP3
PI3K
MEK1/2
Akt
C3G
Rap1
PIP3
p35
Cdc42
Positive
feedback
Strad
ER
CaMKI
AF6
LKB1
RSK
ERK1/2
GSK-3β
Par6
Par3 aPKC
RhoGEF
Cdk5
RhoA
p27kip1
Rho-K
p190
RhoGAP
PAK1
LIMK
Rnd2
Shootin1
Cofilin
Smurf1
UPS
Rac
RacGEF
DCX
SAD-A/B
MAP2/4
JNK1/2/3
Tau1
Microtubule dynamics
SCG10
STEF/Tiam1
CRMP-2
Numb
JIP1
Endocytosis
Slp1
Rab27
Kinesins
Sra-1 WAVE
Transport
MLC
Actin filament dynamics
RP58
Ngn2
Gene expression
Axon initiation & Multipolar-to-bipolar transition
FIG. 6. Signaling pathways involved in neuronal polarization in vivo. See text for details. AC, adenylyl cyclase;
aPKC, atypical protein kinase C; ApoER2, apoE receptor type 2; BDNF, brain-derived neurotrophic factor;
BMP, bone morphogenetic protein; CaMKK, Ca2⫹/calmodulin-dependent protein kinase kinase; Cdc42, cell
division cycle 42; Cdk5, cyclin-dependent kinase 5; CRMP-2, collapsin response mediator protein 2; Dab1,
Disabled-1; DCX, doublecortin; DLK, dual leucine zipper kinase; ER, endoplasmic reticulum; ERK, extracellular
signal-regulated kinase (mitogen-activated protein kinase); GPCR, G protein-coupled receptor; GSK-3␤, glycogen synthase kinase 3␤; IP3, inositol 1,4,5-trisphosphate; JNK, c-jun NH2-terminal kinase; LIMK, LIM domain
kinase; LKB1, liver kinase B1; Lyn, v-yes-1 Yamaguchi sarcoma viral related oncogene homolog; MAP2/4,
microtubule-associated protein 2/4; MARK2, microtubule affinity-regulating kinase 2; MEK, MAPK/ERK
kinase 1; MKK, mitogen-activated protein kinase kinase; MLC, myosin light chain; Ngn2, neurogenin 2; NP-1,
neuropilin-1; NT-3, neurotrophin-3; PAK, p21 activated kinase, Par3, partitioning defective 3; Par6, partitioning defective 6; PIP3, phosphatidylinositol 3,4,5-trisphosphate; PI3K, phosphatidylinositol 3-kinase; PKA,
cAMP-dependent protein kinase; PKG, cGMP-dependent protein kinase; PLC, phospholipase C; p27Kip1,
cyclin-dependent kinase inhibitor 1B; p35, cyclin-dependent kinase 5, regulatory subunit 1; Rho-K, Rho-kinase;
RSK, ribosomal S6 kinase; SAD-A/B, synapses of the amphid-defective kinase-A/B; SCG10, superior cervical
ganglion 10; Sema-3A, semaphorin 3A; sGC, soluble guanylate cyclase; Smurf1, SMAD specific E3 ubiquitin
protein ligase 1; Sra-1, specifically Rac1-associated protein; STEF, Sif and Tiam1-like exchange factor; Strad,
STE20-related kinase adaptor; TAG-1, transient axonal glycoprotein-1; TGF-␤R, transforming growth factor-␤
receptor; Tiam1, T-cell lymphoma invasion and metastasis 1; TrkR, neurotrophic tyrosine kinase receptor;
UPS, ubiquitin-proteasome system; VLDLR, very-low-density lipoprotein receptor; WAVE, Wiskott-Aldrich
syndrome protein (WASP)-family verprolin homologous protein 1.
neuronal polarity are milder than in DN Rac1-expression
studies (191). In contrast, cerebellar granule neurons do not
express other Rac isoforms. The cerebellar granule neurons
of Rac1-deficient mice display impaired neuronal migration
and axon formation both in vivo and in vitro (330). These
phenotypes might be caused by the mislocalization of the
WAVE complex and the subsequently reduced actin dynamics.
Cdc42 plays a critical role in axon formation upstream of
the Par3-Par6-aPKC complex in cultured neurons (298)
and in vivo (93). PI3K activates Cdc42 by producing PIP3,
Cdc42 interacts with Par6 (156, 157, 195, 273), and Par3
mediates Cdc42-induced Rac activation via the specific Rac
GEFs Tiam1 and STEF/Tiam2 (115, 245). Because acti-
vated Rac can bind to PI3K, this mechanism is thought to be
a positive-feedback loop for axon specification.
RhoA is generally associated with actin cytoskeleton remodeling and myosin-based contractility. In neurons,
RhoA usually acts as an inhibitory molecule for the formation of axons and neurites. The CA form of RhoA inhibits
neurite formation, whereas a DN form of RhoA enhances
neurite outgrowth (53, 298). Inhibiting the activity of Rhokinase (ROCK), an effector of RhoA, induces the formation
of multiple axons in cultured neurons (49, 53). Rho-kinase
phosphorylates myosin light chain, myosin phosphatase,
CRMP-2, Tau, MAP2, and LIM kinase and regulates actin
filament and microtubule dynamics and protein trafficking
(6, 7, 13, 173, 201).
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MARK2
NEURONAL POLARITY SIGNALING
Ras, one of the most common oncogenes in humans, is
known to regulate axon specification in cultured neurons
(252, 377). Ras activity is particularly high in the growth
cones of nascent axons (75). Ras activation induces membrane targeting of integrin-linked kinase (ILK), which, in
turn, activates Akt and inactivates GSK-3␤, thereby stimulating axon formation. Because ILK is a pseudokinase, ILK
may act as a scaffold for Ras/Akt/GSK-3␤ (272). Ras also
activates ERK2 through Raf and MEK (9, 62, 180, 185,
236, 263, 355), thereby regulating neuronal polarization
(88, 252, 377).
I) Ca2⫹/calmodulin-dependent protein kinase kinases, CaMKK/CaMKI. Calcium is known to be involved in
axon formation by mediating responses to axon guidance
molecules. The Ca2⫹ effector CaMKK and its target
CaMKI have been implicated in axon elongation in cultured
hippocampal and cortical neurons (55, 351, 362). CaMKI
phosphorylates and activates MARK2 and promotes neurite outgrowth (351). A rapid increase in Ca2⫹ induced by
NT-3 in an inositol 1,4,5-trisphosphate (IP3)-dependent
manner leads to the local activation of CaMKK in the
growth cone (233). Inhibiting neurotrophin receptors or
CaMKK attenuated NT-3-induced axon specification in
cultured neurons and the MP-to-BP transition in vivo (233).
Another report showed that treatment with muscimol, a
GABAA receptor agonist, promoted axonal growth via the
CaMKK/CaMKI pathway, suggesting that distinct upstream factors may regulate CAMKK activity under different circumstances (1).
B) KINASES.
II) The mitogen-activated protein kinase (MAPK) family.
The MEK-ERK2 pathway and the upstream molecule
Ras regulate neuronal polarization (88, 252, 377). The
overexpression of dominant-negative or constitutively
active mutants of MEK1 inhibits proper neuronal polarization (88), suggesting that the spatial and temporal
regulation of the MEK-ERK2 pathway is critical for the
neuronal polarization. ERK2 is localized not only in
the cell soma but also in the growth cone, and it regulates
the interaction between Par3 and KIF3A as described
above (88).
1008
C) SCAFFOLD PROTEINS.
Several important regulators of neuronal polarity play roles as scaffold proteins. In this review,
we focus on the major scaffold proteins for neuronal polarity, CRMP-2 and Par3.
I) CRMP-2. CRMP-2 is one of at least five CRMP isoforms
and historically was independently identified as Ulip2/
CRMP-62/TOAD-64/DRP-2 (102). CRMP-2 is highly expressed in the developing nervous system. Mutations in
UNC-33, a C. elegans homolog of CRMPs, lead to severely
uncoordinated movement and abnormalities in axon guidance (121, 348). The functions of CRMP-2 in neuronal
polarization have been well-studied using cultured neurons.
Previous studies showed that CRMP-2 is enriched in the
growing axons of hippocampal neurons and that CRMP-2
overexpression induces primary axon elongation and multiple axon formation, whereas inhibiting CRMP-2 function
impairs axon formation in cultured hippocampal neurons
(148). CRMP-2 interacts with tubulin heterodimers and
promotes microtubule assembly in vitro (85). CRMP-2 also
regulates the endocytosis of specific adhesion molecules,
including L1, by interacting with Numb (243) and by reorganizing actin filaments via the Sra-1/WAVE1 complex
(166), and it induces the trafficking of vesicles containing
TrkB by interacting with the Slp1/Rab27 complex (12).
Thus CRMP-2 promotes neurite elongation and axon specification by regulating microtubule assembly, adhesion
molecule endocytosis, actin filament reorganization, and
axonal protein trafficking.
CRMP-2 activity is regulated by COOH-terminal phosphorylation. GSK-3␤ phosphorylates CRMP-2 at Thr-514
and Ser-518 following the priming phosphorylation at Ser522 by cyclin-dependent kinase 5 (Cdk5), thereby inactivating the binding of CRMP-2 to tubulin dimers (378). In
cultured hippocampal neurons, the nonphosphorylated
CRMP-2 level is enriched in axon growth cones. The expression of a CA form of GSK-3␤ inhibits axon formation,
whereas the nonphosphorylatable form of CRMP-2 coun-
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Rap1, a member of the Ras superfamily of GTPases, is also
required for appropriate neuronal polarity (298). Rap1 activity is highest in the longest neurite (232). Neurons deficient in the Rap1 GEF C3G (Rapgef1, Grf2) display MP
morphology and are arrested in migration in brain slice
cultures (357). A recent study demonstrated that Reelin, an
extracellular matrix glycoprotein that helps to regulate the
processes of neuronal migration and positioning during the
organization of the cortex, stimulates the activation of
Rap1, regulating the polarity of migrating neurons in the
developing mouse cortex via the regulation of N-cadherin
(81, 158).
JNK, a member of the MAPK family that was initially described as a stress-activated protein kinase, was identified
based on its ability to phosphorylate c-Jun (186). JNK
phosphorylates microtubule regulators, such as DCX,
MAP1B, MAP2, and SCG10/stathmin-2. Activation of
JNK and the JNK binding partner JIP is required for axon
initiation in vitro (54, 254). JNK activity is elevated in the
IZ during cortical development (337). The targeted deletion
of the JNK activator mkk4 (358) or the electroporation of
neurons with dominant-negative JNK inhibits radial migration (167). Recent studies showed that JNK regulates the
MP-to-BP transition and neuronal migration via SCG10/
stathmin-2 phosphorylation during cortical development
(337, 365). JNK1 and dual leucine zipper kinase (DLK)
double-knockout mice exhibited severe defects in axon formation in the cerebral cortex (129).
NAMBA ET AL.
teracts the inhibitory effects of GSK-3␤, suggesting that
GSK-3␤ regulates neuronal polarity via phosphorylating
CRMP-2. Neurotrophic factors, including NT3 and BDNF,
inhibit GSK-3␤ activity and, therefore, CRMP-2 phosphorylation. Depleting CRMP-2 prevents NT3-induced axon
outgrowth in cultured hippocampal neurons (378). Taken
together, regulating GSK-3␤ and CRMP-2 activities is important for neuronal polarization in vitro. The details of the
upstream GSK-3␤ signaling pathway are presented below
(see cell extrinsic signals).
II) Par3. The Par genes were initially identified via genetic
screens for regulators of cytoplasmic partitioning in early
embryos of C. elegans (170). Six par genes were identified
based on this screen. Par1 and Par4 encode serine threonine
kinases, as described above (109, 227). Par2 contains a
RING finger domain that may function in the ubiquitination pathway (218). Par3 and Par6 contain PDZ domains,
suggesting that these proteins act as scaffold proteins (72,
361). Par5 is a member of the 14-3-3 family of proteins,
which bind to phosphorylated serine and threonine residues
(228).
Par3 forms a conserved protein complex with Par6 and
atypical protein kinase C (aPKC), which is defined as the
Par complex. The Par complex regulates multiple polarity
events in animals, including asymmetric cell division, tight
junction formation, and directional migration (67, 100,
152, 176, 325). During neuronal development, the Par
complex is mainly localized at the tips of nascent axons in a
KIF3A-dependent manner, and it participates in axon formation (244, 305), implying that the transport of the Par
complex by kinesin plays an important role in axon formation, as shown in hippocampal neurons (244). In this process, Par3 acts as a cargo adaptor by linking KIF3A with the
The Par complex is a key factor in the Rac1/Cdc42 signaling
pathway. Cdc42 plays a critical role in axon formation
upstream of the Par complex (298). Activated Cdc42 interacts with Par6 via the semi-Cdc42/Rac interactive binding
(CRIB) domain of Par6, which then associates with aPKC
and Par3, leading to aPKC activation (156, 157, 195, 273).
Par3 directly interacts with Tiam1/STEF, the Rac-specific
GEFs, and forms a complex with aPKC, Par6, and Cdc42,
thereby mediating Cdc42-induced Rac1 activation (211,
245). Because Cdc42 is activated by PI3K and the active
form of Rac1 can activate PI3K, a positive-feedback loop
composed of PI3K, Cdc42, the Par complex, and Rac1 has
been considered responsible for neuronal polarization. This
pathway promotes axon formation, presumably via actin
filament remodeling. In addition, Par3 acts as a microtubule-associated protein that binds to and bundles microtubules in a conformation-dependent manner (47). In the
closed conformation, the intramolecular interaction between the NH2 and COOH termini of Par3 impairs its
microtubule regulatory activity. As Par3 accumulates, intermolecular oligomerization promotes an open conformation that directly binds to, bundles, and stabilizes microtubules via its NH2 terminus. This microtubule regulatory
activity of Par3 is crucial for neuronal polarization (47).
B. Signals That Regulate Neuronal
Polarization In Vivo
Polarization of neurons in vivo is precisely regulated by
their microenvironment (extracellular signals) and the timing of gene expression. The extracellular molecules, including secreted factors, such as neurotrophins, TGF-␤, Wnts,
IGF-I, semaphorin, and Reelin; ECM components, such as
laminin; and cell adhesion molecules, such as TAG-1 and
N-cadherin, are known to regulate neuronal polarization in
cultured neurons and also in vivo (81, 92, 127, 158, 233,
237, 269, 376) (FIGURE 3). Of these, we first review neurotrophins, TGF-␤, Reelin, TAG-1, and N-cadherin, which
are known to regulate neuronal polarization in vivo. Generally, these extracellular signals use similar intracellular
signal molecules, such as CaMK, LKB1, and Rac1, to induce neuronal polarization. In some cases, extracellular signals such as semaphorins antagonize the intracellular signal
pathways that are known to initiate axon formation in cul-
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The function of CRMP-2 in neuronal polarization in vivo
has recently been analyzed. In the developing neocortex,
inhibiting CRMP-2 function via gene knockdown or via the
expression of a dominant-negative mutant delays neuronal
migration and impairs the MP-to-BP transition (324). A
recent study demonstrated that the phosphorylation status
of CRMP-2 affects the MP-to-BP transition (151). There
are few in vivo studies of CRMP-2 using knockout mice. In
contrast to the studies using cultured neurons, there are no
apparent axonal abnormalities in knock-in mice expressing
the phosphorylation-deficient form of CRMP-2 (CRMP-2S522A) (374). However, these knock-in mice exhibit increased numbers of primary dendrites. This phenotype is
enhanced by crossing with the CRMP-1-knockout mice
(374). Because neurons express all CRMPs (CRMP-1
through CRMP-5) during brain development, functional
compensation may occur in individual CRMP knockout
models. Further studies using individual and multiple conventional or conditional CRMP knockout mice are required.
Par complex. A recent study demonstrated that the Par3KIF3A interaction is regulated by the ERK2-dependent
phosphorylation of Par3 and is important for neuronal polarization (88). More recently, two studies showed that
Par3 regulates neuronal polarization in vivo. In the developing cerebral cortex, knockdown of Par3 impairs the MPto-BP transition (47, 88). Interestingly, unlike WT Par3,
mutant Par3 that does not interact with KIF3A fails to
rescue the knockdown phenotype, suggesting that Par3mediated protein transport is also important for neuronal
polarization in vivo (88).
NEURONAL POLARITY SIGNALING
ture and reciprocally induce dendrite formation (303). Thus
the key components that drive the polarization machinery
are shared by neurons in culture and in vivo. However, the
extracellular molecules that trigger the shared intracellular
molecules are diverse among the developmental stages and
types of neurons. This diversity generates redundancy and
may ensure proper neuronal polarization in vivo.
Given that transcription factors and epigenetic modifiers
determine the timing of neuronal differentiation and morphogenesis (206), we next review how the regulation of
gene expression contributes to neuronal polarization in
vivo.
1. Extracellular signals
Neurotrophins also activate LKB1 through cAMP and PKA
or p90RSK activation (302). LKB1, a mammalian ortholog
of the serine/threonine kinase Par4 (107), is known to regulate cell polarity in various cell types (16). Emx-Cre-mediated conditional knockout of the LKB1 gene in mice leads
to the absence of axons from cortical pyramidal neurons
without affecting their migration (28). Knockdown of
LKB1 prevents axon specification both in cultured neurons
1010
B) TGF-␤ SUPERFAMILY PROTEINS. The TGF-␤ superfamily consists of TGF-␤, activin, and bone morphogenetic protein
(BMP) and regulates a wide variety of cellular processes.
Upon ligand binding, the TGF-␤ superfamily receptors
form heterotetrameric complexes composed of type I and
type II serine/threonine kinases. In type II TGF-␤ receptor
(Tgfbr2) conditional knockout mice, neuronal migration is
delayed, and axons fail to form in the developing neocortex
(376). Tgfbr2 phosphorylates Par6 at S345 (376). Depending on this phosphorylation, Smurf1, a ubiquitin ligase, is
recruited to the activation site and degrades RhoA (49,
257), suggesting that TGF-␤ signaling can modulate the
local stoichiometry of Rho GTPases.
Because TGF-␤2 expression is restricted to the VZ, there
might be a gradient of TGF-␤2 concentration along the
ventricular-to-pial-surface axis (376). However, this simple
radial gradient cannot explain the direction of axon initialization because more than half of the MP cells initially
extend axons tangentially rather than radially toward the
ventricle (119, 237, 287). Therefore, the TGF-␤ gradient
likely acts as a basal factor for axon specification, and other
cues, such as cell adhesion molecules, may select one immature
neurite as an axon, as described below (237). Another TGF-␤
superfamily member, BMP, suppresses CRMP-2 transcription
via SMAD during brain development (324). Therefore, there
are two TGF-␤ signaling pathways that regulate neuronal
polarization: the Par6/Smurf1-mediated local protein deg-
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A) NEUROTROPHINS. The neurotrophins consist of four members: nerve growth factor (NGF), BDNF, NT-3, and neurotrophin-4 (NT-4). Among them, BDNF and NT-3 are
highly expressed in the central nervous system and stimulate axon specification and axon elongation (221, 233,
302). Neurotrophins activate specific receptor tropomyosin
receptor kinases (Trks) and induce Ras and phosphoinositide 3-kinase (PI3K) activation, thereby increasing the
production of phosphatidylinositol 3,4,5-trisphosphate
(PIP3) (281). PIP3 is enriched at the tips of nascent axons,
and treatment of cultured neurons with PI3K inhibitors
disturbs axon specification (210, 305). PIP3 indirectly
activates Akt, thereby inactivating GSK-3␤ via its phosphorylation at serine 9. GSK-3␤ inactivation leads to the
activation of microtubule assembly-promoting proteins,
such as Tau1 and CRMP-2 (154, 378). In addition, neurotrophins activate CaMKK and CaMKI through Ca2⫹ induced by IP3-induced calcium release (IICR) as described
above (233). CaMKI can activate Par1/MARK2. Par1 is a
serine/threonine kinase (109), and its mammalian ortholog,
microtubule affinity-regulating kinase (MARK), was originally identified as a modulator of microtubule dynamics
(69). MARK2 phosphorylates MAPs, including tau,
MAP2/4, Tau1, and DCX, and changes their affinity to
microtubules (31, 69, 293). MARK2 is negatively regulated
by aPKC-dependent phosphorylation (48). Neurons electroporated with MARK2 or DCX shRNA remain in the IZ
and exhibit an MP morphology in vivo (291, 292). Given
that increasing or decreasing the expression of MARK2
alters neuronal migration and the MP-to-BP transition, the
appropriate regulation of MARK2 expression is essential
for neuronal polarization in vivo (291).
and in vivo (302) and prevents neuronal migration in vivo
(13). LKB1 activity is regulated by cAMP and PKA (301,
302). In contrast, semaphorin-3A (Sema3A), a secreted
chemorepulsive protein, elevates the cGMP level but reduces the cAMP level reciprocally, thereby suppressing the
PKA-dependent phosphorylation of LKB1 (303). Once
LKB1 is activated via PKA or p90RSK-dependent phosphorylation, LKB1 interacts with the pseudokinase STRAD
and phosphorylates several downstream kinases, such as
NUAK, AMPK MARK2, and the SAD kinases (197). Of
these kinases, SAD kinases and MARK2 likely mediate the
function of LKB1 in neuronal polarization. SAD kinases,
which belong to the MARK family, phosphorylate Tau-1 at
S262, which is highly phosphorylated in dendrites but not
in axons (173). SAD-A and SAD-B kinase double-knockout
mice exhibited abnormally oriented dendrites and a loss
of axons in vivo (173). Suppression of other kinases, such
as NUAK and AMPK, did not affect neuronal polarization (52, 366), but overactivation of AMPK decreased
the axon length in vivo (8). Notably, NUAK regulates
terminal axon branching both in cultured neurons and in
vivo (52). Taken together, the neurotrophins precisely regulate the activity of several protein kinases. However, these
kinases may not simply work in parallel. Therefore, it will
be important to show when and where each of these molecules works during the polarization process.
NAMBA ET AL.
radation of RhoA and the SMAD-dependent transcriptional regulation of CRMP-2 expression.
C) REELIN.
The effect of Reelin on axon formation has been examined
using cultured neurons but remains controversial. One
study showed that stimulating cultured neurons with fulllength Reelin increased growth cone motility and induced
axon formation via Cdc42 activation (189). Another study
showed that stimulating one minor neurite with the cleaved
form of Reelin did not induce axon formation, unlike other
axon formation-inducing factors, such as neurotrophins,
TGF-␤, and Wnts (233). These results raise the possibility
that the status of posttranslational processing is important
for its function in axon formation, as described previously
(159).
D) CELL ADHESION MOLECULES. I) TAG-1. TAG-1 is a GPIanchored immunoglobulin superfamily cell adhesion molecule that is known to regulate axon fasciculation and guidance (163, 308). TAG-1 protein expression is highest in the
lower IZ, where the MP-to-BP transition and axon specification occur (237). In the lower IZ, TAG-1 is expressed
in both MP cells and pioneering axons. TAG-1 can mediate cell-to-cell interactions between these cells. Suppression of TAG-1 expression in MP cells by shRNA
disrupts the MP-to-BP transition and axon specification.
WT TAG-1 rescued these phenotypes, but the cell adhesion-deficient mutants of TAG-1 did not (237).
II) N-cadherin. N-cadherin is a member of the classical
cadherin family that mediates Ca2⫹-dependent adhesion
(130). In the embryonic cerebral cortex, N-cadherin is expressed in both radial glial cells and neurons and mediates
the interaction between these two cell types (81, 91, 120,
158, 168). N-cadherin-mediated cell-to-cell interactions are
important for radial glia-guided neuronal migration. A recent study demonstrated that this type of migration is precisely regulated by N-cadherin trafficking in neurons (168).
Inhibition of the endocytic or recycling process via transfection with Rab5 or Rab11 dominant-negative mutant, respectively, reduces N-cadherin surface expression in neurons, thereby affecting neuronal migration. Although the
surface expression of N-cadherin has been reported to be
regulated by Reelin signaling via Rap1, as described above
(81, 158), the mechanism by which Reelin signaling regulates Rab5- and Rab11-dependent trafficking remains unknown. Therefore, the functional interactions between
Rap1 and Rab5 or Rab11 must be elucidated.
N-cadherin is also involved in the MP-to-BP transition in
cultured neurons (92). Nonpolarized stage 1 neurons
generate their initial neurites from the N-cadherin-contacting site. In the embryonic cerebral cortex, N-cadherin
has been shown to regulate the orientation of the leading
process of BP cells (92). In addition, inhibiting N-cadherin function using a dominant-negative mutant impairs
the MP-to-BP transition (81, 91, 158, 168). Taken together, these results indicate that N-cadherin regulates
leading process formation during the MP-to-BP transi-
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Reelin is a glycoprotein that is secreted by CajalRetzius cells into the marginal zone of the embryonic cerebral cortex (341). Because cortical layer formation is severely affected in Reelin-deficient mice, this molecule is
thought to be an important regulator of cortical development. Reelin acts through its receptor, apolipoprotein E
receptor 2 (ApoER2), as well as very-low-density lipoprotein receptor (VLDLR), and activates several intracellular
signaling molecules via the adaptor protein disabled-1
(Dab1). Recent studies suggested that Reelin signaling regulates the MP-to-BP transition and axon formation (81,
158). Reelin stimulation induces Dab1 phosphorylation
and the binding of Crk-family proteins (CrkL) and C3G, a
Rap1GEF, to Dab1 (24). This complex activates Rap1 in
a Reelin-dependent manner. Dab1- or Rap1-deficient neurons exhibited MP morphology in vivo, suggesting that
Reelin-Rap1 signaling is important for the MP-to-BP transition (81, 158). A previous study suggested that Rap1 activation was required for the appropriate targeting of cadherin proteins to maturing cell-cell interactions (134). Indeed, Rap1 regulates the surface expression of N-cadherin
in neurons (158), possibly through p120 catenin (142). Because N-cadherin has been shown to be important for the
MP-to-BP transition and migration (81, 91, 158, 168), the
function of Reelin in these events may be partially mediated
by N-cadherin. The detailed roles of N-cadherin are described in the following section.
As with other GPI-anchored proteins, TAG-1 can indirectly activate intracellular signal molecules (163, 308).
The outside-in signaling of TAG-1 is mediated by two
possible mechanisms: one involving transmembrane proteins that associate with TAG-1 in a cis manner and
another involving Src family kinases (SFKs) localized to
lipid rafts (163). Previous studies identified several transmembrane proteins, including L1, NrCAM, and contactin-associated protein-like 2 (CASPR2), as cis-associating proteins for TAG-1. Although these molecules have
been shown to be involved in neuronal migration (175,
264, 342), they seem not to be involved in the TAG-1mediated MP-to-BP transition or axon specification in
vivo (237). TAG-1 colocalizes with SFKs, including Src,
Fyn, and Lyn, in lipid rafts (164, 165, 270). Among these
SFKs, Lyn is a downstream kinase of TAG-1 in cultured
neurons (164, 165) and in vivo (237), and it regulates
neuronal polarization (237). Given that Lyn can activate
Rac1 (58), the TAG-1-Lyn signaling pathway might activate Rac1 in neurons to induce polarization (237) (FIGURE 3). In conclusion, TAG-1-dependent cell-to-cell interactions between MP cells and pioneering axons activate Lyn and Rac1 and promote neuronal polarization
(237).
NEURONAL POLARITY SIGNALING
tion and subsequently mediates radial glial cell-dependent neuronal migration.
2. Regulation of gene expression
The members of the forkhead transcription factor FoxO
family (FoxO1, 3 and 6) and FoxG1 regulate neuronal migration, the MP-to-BP transition, and axon-dendrite specification in cultured neurons and in vivo (57, 215). FoxO
knockdown reduces the expression of Par6, Pak1, R-Ras,
APC, and CRMP-2 (57). Among these genes, the Pak1 promoter contains putative FOXO-binding sites; in fact, Pak1
transcription is directly regulated by FoxO (57). FoxG1
knockout mice show altered expression of several genes
that may be involved in neural development (215). The
most strongly downregulated gene is disabled homolog 1
(Dab1), a key factor in Reelin signaling. The most strongly
1012
B) EPIGENETIC MODIFIERS. Studies of epigenetics have advanced
remarkably during the past decade. Epigenetic mechanisms,
such as DNA methylation, histone modification, and
miRNA, are important for the expression of genes that
regulate neural development (128). Most epigenetic studies
have focused on progenitor cell differentiation and cell-type
specification; however, several studies have demonstrated
the epigenetic control of neuronal polarization. miRNAs
are produced from pre-miRNA following processing mediated by the RNAse III enzyme Dicer. Neuronal differentiation, dendrite morphogenesis, and axon formation are affected in Dicer knockout mice (194, 356). In addition, cultured hippocampal neurons from Dicer knockout mice
exhibit increased numbers of neurons displaying multiple
axons (194). Various types of miRNAs regulate the expression of polarity-regulating genes. miR-219 regulates the expression of Par3 and aPKC in zebrafish larvae (147). miR134 inhibits neuronal migration in the embryonic cerebral
cortex by suppressing Dcx gene expression (94). miR-124
regulates the expression of the corepressor for element-1silencing transcription factor (CoREST) in Xenopus retinal
ganglion cells (28) and in mouse (356). miR-22 also regulates the expression of CoREST (356). CoREST forms a
complex with REST and represses target gene expression by
recruiting chromatin-binding proteins and histone modifiers (274). In the developing cerebral cortex, CoREST and
REST regulate Dcx gene expression, thereby regulating
neuronal migration (84, 202). Epigenetic regulation of polarity genes may fine-tune gene expression in a time- and
space-dependent manner. Interestingly, miRNA can be
transported to the extracellular space (321). Therefore,
miRNA may act as a microenvironmental cue for neuronal
development.
IV. THEORETICAL BASIS OF NEURONAL
POLARIZATION
Various molecules have been identified as regulators of
neuronal polarity, as described above. However, integrating and assembling these signaling molecules into
one model might be important for understanding the
entire picture of neuronal polarity. One suitable method
is mathematical modeling (149, 239, 349). Under physiological conditions, neurons generally extend one axon,
and other minor neurites never generate an axon. How is
this winner-takes-all process accomplished? A local activation-global inhibition mechanism is a likely model that
explains this winner-takes-all process (11, 149, 239).
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A) TRANSCRIPTION FACTORS. Proneural transcription factors
are known to be important regulators of neural development (108). Conventionally, these molecules are primarily
considered to be involved in cell-type specification. Recent
studies have suggested that basic helix-loop-helix (bHLH)
proneural transcription factors regulate not only neurogenic differentiation but also neuronal migration and the
MP-to-BP transition (125, 258). In the embryonic cerebral
cortex, neurogenin2 (Ngn2) induces neuronal migration
and the MP-to-BP transition by negatively regulating
RhoA, presumably via two different mechanisms (95, 117,
241). Ngn2 negatively regulates RhoA transcription. In
contrast, Ngn2 positively regulates Rnd2 transcription. The
Rnd subfamily (Rnd1, Rnd2, and Rnd3/RhoE) represents a
new branch of the Rho family of GTPases (78, 105, 247).
Unlike other Rho family members, Rnd proteins lack GTPase
activity. Thus they have been considered constitutively active (78, 105, 247), and they are thought to be regulated
primarily at the level of their expression (46). Rnd2 activates p190RhoGAP, a GAP of RhoA, thereby inhibiting
RhoA activity (364). Rnd2-deficient neurons display MP
morphology with long processes, suggesting that Rnd2 is
important for the MP-to-BP transition (126, 236). Rnd2
may be involved in the trafficking of membrane-bound molecules that regulate the MP-to-BP transition (126, 265).
Rnd3 induces F-actin depolymerization, resulting in the stabilization and attachment of the leading process to a radial
glial fiber via the regulation of RhoA activity (265). Furthermore, the zinc-finger transcriptional repressor RP58
and chicken ovalbumin upstream promoter-transcription
factor I (COUP-TFI), an orphan nuclear receptor, also repress Rnd2 expression and regulate neuronal migration and
the MP-to-BP transition (126, 251). Ngn2 also positively
regulates the expression of Dcx and p35 (95), both of which
are known to be regulators of neuronal migration and the
MP-to-BP transition. Interestingly, p35 activates CDK5
and then stabilizes the Ngn2 protein via the CDK inhibitor
p27 (Kip1) (241). Therefore, this positive feedback system
represses RhoA activity and expression.
upregulated gene is deleted in autism 1 (DIA1, GoPro49).
DIA1 is localized to the Golgi complex and is primarily
expressed in mesenchymal and cartilaginous tissues but is
also mildly expressed in the cortical hem of the cerebral
cortex (332). Although this gene has been implicated in
autism spectrum disorder (ASD), its function remains unclear (15, 226).
NAMBA ET AL.
A
Positive signal
Positive
feedback signal
Negative
feedback signal
B
Activity of factor Y (%)
100
State: Axon
80
60
tration of factor X and the activity of factor Y act as a
bistable switch. When the concentration of factor X exceeds the threshold, then factor Y is in the “on state.”
Once factor Y is activated, its activity is sustained even if
the concentration of factor X is reduced. Thus the specific
neurite continuously elongates and ultimately develops
into an axon. In contrast, because the concentration of
factor X depends on the length of the neurite, the concentration of factor X in other neurites cannot increase,
and these neurites fail to elongate. Notably, negative
feedback signaling molecules are not necessary in this
model. Whether the negative feedback signal is a retrograde propagation of the signaling molecules or a lower
concentration of factor X in the soma and in other minor
neurites remains controversial. Experimentally identifying the negative feedback signal is a critical issue regarding neuronal polarity. The possible candidates include
second messengers. It has been proposed that cAMP and
cGMP have antagonistic effects on each other (304). Locally induced cAMP elevation induces PKA activation
and subsequent axon initiation. Notably, the cAMP level
in the unstimulated neurites begins to decline just after
the stimulation, and the cGMP level in these neurites is
reciprocally increased. This result suggests that cAMP
acts as a long-range negative feedback signal. However,
it remains unclear whether cAMP directly transmits the
negative signal from the stimulated neurite to other neurites. One of the axon-initiating extracellular molecules,
neurotrophin, induces not only cAMP elevation but also
IICR or Ras activation (233, 302), suggesting that these
molecules potentially transmit the long-range negative
signals. It has also been shown that cGMP stimulation
does not reduce the cAMP level in other neurites (304).
Taken together, these results suggest that the positive
signal (cAMP) can generate a long-range negative signal
for other neurites but that the negative signal (cGMP)
cannot induce long-range positive signals. Therefore,
positive stimulation (e.g., neurotrophin) seems to be
more critical for the timing of neuronal polarization than
negative stimulation (e.g., semaphorin).
40
20
State: Minor neurite
0
The concentration of X at tips of neurites
FIG. 7. Theoretical model for neuronal polarization. A: a local
activation-global inhibition mechanism using positive- and negativefeedback signals. B: bistable switch for neuronal polarization. The
selective accumulation of positive factors (termed factor X) is regulated by the active anterograde transport, diffusion, and degradation
of these positive factors. The activity of the signal molecule (termed
factor Y) that induces the neurite elongation depends on the concentration of factor X in a hysteretic manner. Therefore, the concentration of factor X and the activity of factor Y act as a bistable
switch. When the concentration of factor X exceeds the threshold,
then factor Y is in the “on state.” Once factor Y is activated, its
activity is sustained even if the concentration of factor X is reduced.
Thus the neurite stays in the “axon state” (magenta).
Furthermore, the local accumulation and activation of
factor X drive a positive feedback loop. Several studies
have experimentally demonstrated that the Par complexRac1-Cdc42 and HRas-PI3K positive feedback loops induce axon formation (11). In addition, turning off the
negative feedback loops might be a positive feedback
signal. The GEF-H1-RhoA-Rho-kinase axis may act as a
negative feedback loop. Once Rho-kinase is activated by
GEF-H1 and RhoA, Rho-kinase phosphorylates and inactivates p190RhoGAP (224). This process might lead to
sustained RhoA activation. Because suppression of
GEF-H1 and Rho-kinase expression in cultured neurons
induces the formation of multiple axons (49, 51, 53), this
pathway plays critical roles in the “stochastic” neuronal
polarization. PKA activates LKB1 and thereby induces
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This model contains three key components: a bistable
switch, a negative feedback signal(s), and a positive feedback signal(s) (240, 344) (FIGURE 7). The selective accumulation of positive factors (termed factor X), such as
CRMP-2 (for microtubule polymerization and receptor
trafficking), Par complex (for Rac1 activation), and
HRas and Shootin1 (for PI3K activation), has been
shown to facilitate the development of a neurite into an
axon (148, 244, 345). This accumulation is regulated by
the active anterograde transport, diffusion, and degradation of these positive factors. Because anterograde and
retrograde transport are primarily mediated by motor
proteins and by diffusion, respectively, positive factors
might accumulate at the tips of the longest neurites. If the
rate of motor protein-mediated transport is equivalent to
or lower than that of diffusion, it is difficult for neurons
to establish axons in the simulation (240, 344). Indeed,
inhibition of Kinesin-1 function in neurons abolished
axon formation (174). Then, the local accumulation of
factor X activates signaling molecules (termed factor Y,
e.g., PIP3, cAMP, and Ca2⫹) that induce neurite elongation via downstream pathways as described in section III.
The activity of factor Y depends on the concentration of
factor X in a hysteretic manner. Therefore, the concen-
NEURONAL POLARITY SIGNALING
axon initiation, as described above. At the same time,
PKA can inhibit the activation of RhoA through the inactivation of GEF-H1 (209). Therefore, PKA activation
drives neuronal polarization in two different ways: activation of a positive signal and inhibition of a negative
feedback loop. Extracellular signals such as neurotrophin and cell adhesion might trigger the positive feedback loops.
V. MAINTENANCE OF NEURONAL
POLARITY
1014
Several genes that regulate neuronal polarity have also
been identified as causal genes of neurodevelopmental
diseases. Of these diseases, we focus on lissencephaly and
ASD. Lissencephaly is a malformation of cerebral cortical development and is categorized into group IIB (26).
This malformation is due to generalized abnormal neuron migration. Because the neurons migrate improperly,
the brain becomes smooth, lacking most of the typical
sulci and gyri. Thus far, several gene mutations have been
found in lissencephalic patients. Most of these mutations
are associated with microtubule dynamics or Reelin signaling. In the early 1990s, Ledbetter and colleagues (65,
188) identified microdeletions in 17q13.3 of patients
with Miller-Dieker syndrome, a type 1 lissencephaly (64,
188). Then, Reiner et al. (282) identified a causal gene,
termed LIS1 (282). Other groups identified the DCX
gene as a causal gene of X-linked subcortical laminar
heterotopia (SCLH) (60, 98). Mutations in TUBA1A
(previously referred to as TUBA3) were identified in a
patient with lissencephaly without any mutations in the
LIS1 or DCX gene (169, 182, 268). All of these genes,
which correspond to tubulin or to microtubule modulators, have been experimentally demonstrated to regulate
neuronal migration, as described above. Another causal
gene of lissencephaly is associated with Reelin signaling.
Mutations in RELN or VLDLR were found in patients
with autosomal recessive lissencephaly (36, 137). Based
on the finding that the corpus callosum is thin in these
patients, Reelin signaling regulates not only neuronal migration but also axon formation during human brain
development, which is identical to its role in rodents.
Although the etiology of ASD remains unknown, many
researchers tend to understand the etiology of ASD in
terms of neurodevelopmental disabilities (220). Several
susceptibility genes of ASD are known to regulate neuronal migration and morphogenesis. In some ASD patients,
several SNPs were identified in the 3’ untranslated region
of MARK1, thereby increasing the expression level of
MARK1 (207). Because precise regulation of MARK2
expression is important for neuronal migration and for
the MP-to-BP transition in the mouse cerebral cortex
(291), neuronal polarity might be affected in these ASD
patients.
The fundamental mechanisms that regulate axon specification may contribute to axon regeneration. For example, microtubule stabilization induces axon regeneration
after nerve injury (124, 299). Axon regeneration after
optic nerve axotomy is enhanced in phosphatase and
tensin homolog (PTEN) knockout mice, as PTEN exerts
an antagonistic effect on PI3K (260). Axon regeneration
in PTEN knockout mice is partially, but not completely,
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After neurons are polarized, neurons must maintain their
polarity to perform their function. Experimental manipulations can alter axon-dendrite polarity either in cultured neurons or in vivo (65, 101, 103, 122). Banker and
colleagues (65, 103) demonstrated that axonal transection induces polarity changes in stage 3 neurons. When
the axon is transected at its distal part, the damaged axon
undergoes regrowth. In contrast, axonal transection at
the proximal part induces axon development in one of
the minor neurites. Similar phenomena were detected
when axonal transection was performed on completely
polarized stage 4 or 5 neurons (101). In completely polarized neurons, axonal transection at the proximal part
causes dendrites to adopt axonal identities. These results
suggest that the proximal part of the axon is crucial for
maintaining axon identity. A specific subcellular structure, termed the AIS, is known to be localized to the most
proximal region of the axon in polarized cells (259). The
AIS is composed of the scaffold protein ankyrinG, ␤IV
spectrin, actin filaments, cell adhesion molecules, and
channels (30, 278, 327). The AIS performs three fundamental functions: regulating action potentials, selecting
axon-directed cargo, and maintaining neuronal polarity
(327). A series of studies revealed that disrupting the AIS
in polarized neurons by knocking out ankyrinG causes
axons to acquire characteristics of dendrites, such as the
formation of spines and synapses (122, 314). In contrast,
the loss of the AIS does not affect the establishment of
neuronal polarity (89). One question remains unanswered: How does the AIS maintain neuronal polarity?
One possible mechanism is the AIS-dependent regulation
of transport. A previous study suggested that cargo selection occurs in the AIS (315). At the AIS, the transport
efficiency of the axon-directed motor-cargo complex is
higher than that of the dendrite-directed motor-cargo
complex (315). Because F-actin is enriched at the AIS
(315) and myosin is required for dendrite-directed transport (192, 315), the exclusion of dendrite-directed cargo
from axons may occur at the AIS in a myosin-dependent
manner (192). Because the AIS is required for maintaining the axon (122, 314), preserving neuronal polarity
may require AIS-regulated selective cargo transport.
VI. INVOLVEMENT OF NEURONAL
POLARITY IN DISEASE AND
TRAUMATIC BRAIN INJURY
NAMBA ET AL.
inhibited by treatment with rapamycin, an inhibitor of
mTOR. Therefore, other downstream signaling molecules, such as Akt and GSK-3␤, might be involved in
axon regeneration, suggesting that microtubule stabilization via GSK-3␤ inactivation may induce axon regeneration.
VII. NEW ERA OF NEURONAL POLARITY
ACKNOWLEDGMENTS
Address for reprint requests and other correspondence:
K. Kaibuchi, Dept. of Cell Pharmacology, Nagoya University Graduate School of Medicine, 65 Tsurumai,
Showa, Nagoya 466-8550, Japan (e-mail: kaibuchi@med.
nagoya-u.ac.jp).
GRANTS
This work was supported by Japan Society for the Promotion of Science (JSPS) KAKENHI Grant 25251021 (to K.
Kaibuchi), Ministry of Education, Culture, Sports, Science
and Technology (MEXT) KAKENHI on Innovative Areas
“Neural Diversity and Neocortical Organization” Grant
23123507 (to K. Kaibuchi), and JSPS Grant-in-Aid for
Young Scientists Grant 25830033 (to T. Namba). Part of
this work is the result of “Bioinformatics of Brain Sciences”
conducted under the Strategic Research Program for Brain
Science (SRPBS) by MEXT (to K. Kaibuchi).
1. Ageta-Ishihara N, Takemoto-Kimura S, Nonaka M, Adachi-Morishima A, Suzuki K,
Kamijo S, Fujii H, Mano T, Blaeser F, Chatila TA, Mizuno H, Hirano T, Tagawa Y,
Okuno H, Bito H. Control of cortical axon elongation by a GABA-driven Ca2⫹/
calmodulin-dependent protein kinase cascade. J Neurosci 29: 13720 –13729, 2009.
2. Ahmad FJ, Yu W, McNally FJ, Baas PW. An essential role for katanin in severing
microtubules in the neuron. J Cell Biol 145: 305–315, 1999.
3. Akella JS, Wloga D, Kim J, Starostina NG, Lyons-Abbott S, Morrissette NS, Dougan
ST, Kipreos ET, Gaertig J. MEC-17 is an alpha-tubulin acetyltransferase. Nature 467:
218 –222, 2010.
4. Akiyama H, Matsu-ura T, Mikoshiba K, Kamiguchi H. Control of neuronal growth
cone navigation by asymmetric inositol 1,4,5-trisphosphate signals. Science Signaling 2:
ra34, 2009.
5. Amann KJ, Pollard TD. Direct real-time observation of actin filament branching mediated by Arp2/3 complex using total internal reflection fluorescence microscopy.
Proc Natl Acad Sci USA 98: 15009 –15013, 2001.
6. Amano M, Ito M, Kimura K, Fukata Y, Chihara K, Nakano T, Matsuura Y, Kaibuchi K.
Phosphorylation and activation of myosin by Rho-associated kinase (Rho-kinase). J Biol
Chem 271: 20246 –20249, 1996.
7. Amano M, Kaneko T, Maeda A, Nakayama M, Ito M, Yamauchi T, Goto H, Fukata Y,
Oshiro N, Shinohara A, Iwamatsu A, Kaibuchi K. Identification of Tau and MAP2 as
novel substrates of Rho-kinase and myosin phosphatase. J Neurochem 87: 780 –790,
2003.
8. Amato S, Liu X, Zheng B, Cantley L, Rakic P, Man HY. AMP-activated protein kinase
regulates neuronal polarization by interfering with PI 3-kinase localization. Science
332: 247–251, 2011.
9. Anderson NG, Maller JL, Tonks NK, Sturgill TW. Requirement for integration of
signals from two distinct phosphorylation pathways for activation of MAP kinase.
Nature 343: 651– 653, 1990.
10. Arimura N, Hattori A, Kimura T, Nakamuta S, Funahashi Y, Hirotsune S, Furuta K,
Urano T, Toyoshima YY, Kaibuchi K. CRMP-2 directly binds to cytoplasmic dynein
and interferes with its activity. J Neurochem 111: 380 –390, 2009.
11. Arimura N, Kaibuchi K. Neuronal polarity: from extracellular signals to intracellular
mechanisms. Nat Rev Neurosci 8: 194 –205, 2007.
12. Arimura N, Kimura T, Nakamuta S, Taya S, Funahashi Y, Hattori A, Shimada A,
Menager C, Kawabata S, Fujii K, Iwamatsu A, Segal RA, Fukuda M, Kaibuchi K. Anterograde transport of TrkB in axons is mediated by direct interaction with Slp1 and
Rab27. Dev Cell 16: 675– 686, 2009.
13. Arimura N, Menager C, Kawano Y, Yoshimura T, Kawabata S, Hattori A, Fukata Y,
Amano M, Goshima Y, Inagaki M, Morone N, Usukura J, Kaibuchi K. Phosphorylation
by Rho kinase regulates CRMP-2 activity in growth cones. Mol Cell Biol 25: 9973–
9984, 2005.
14. Aumais JP, Tunstead JR, McNeil RS, Schaar BT, McConnell SK, Lin SH, Clark GD,
Yu-Lee LY. NudC associates with Lis1 and the dynein motor at the leading pole of
neurons. J Neurosci 21: RC187, 2001.
15. Aziz A, Harrop SP, Bishop NE. Characterization of the deleted in autism 1 protein
family: implications for studying cognitive disorders. PLoS One 6: e14547, 2011.
16. Baas PW, Black MM, Banker GA. Changes in microtubule polarity orientation during
the development of hippocampal neurons in culture. J Cell Biol 109: 3085–3094, 1989.
17. Baas PW, Deitch JS, Black MM, Banker GA. Polarity orientation of microtubules in
hippocampal neurons: uniformity in the axon and nonuniformity in the dendrite. Proc
Natl Acad Sci USA 85: 8335– 8339, 1988.
18. Baas PW, Joshi HC. Gamma-tubulin distribution in the neuron: implications for the
origins of neuritic microtubules. J Cell Biol 119: 171–178, 1992.
DISCLOSURES
19. Baas PW, Lin S. Hooks and comets: the story of microtubule polarity orientation in the
neuron. Dev Neurobiol 71: 403– 418, 2011.
No conflicts of interest, financial or otherwise, are declared
by the authors.
20. Baas PW, Slaughter T, Brown A, Black MM. Microtubule dynamics in axons and
dendrites. J Neurosci Res 30: 134 –153, 1991.
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
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In recent decades, many researchers have focused much
effort on understanding neuronal polarity. As technologies have improved and developed, we have opened the
door and entered new research fields. One of the major
open questions has remained the same for 30 years: how
do neurons generate only one axon? As described above,
a local activation-global inhibition model likely explains
this process; however, this model has not been experimentally verified. One reason for this lack of verification
is the lack of suitable technologies. Given that many signaling molecules, such as Ca2⫹, cAMP, cGMP, and PIP3,
have been shown to mediate neuronal polarization in a
specific subcellular region (4, 233, 303), multiple and
super resolution imaging will aid in the discovery of the
spatiotemporal regulation of these signaling molecules.
In addition, the mathematical prediction of “missing
links” will assist in the discovery and understanding of
key molecules or events. Integrating different fields may
provide additional breakthroughs that could provide further insight into this process.
REFERENCES
NEURONAL POLARITY SIGNALING
21. Bai J, Ramos RL, Ackman JB, Thomas AM, Lee RV, LoTurco JJ. RNAi reveals doublecortin is required for radial migration in rat neocortex. Nat Neurosci 6: 1277–1283,
2003.
22. Baines AJ, Bignone PA, King MD, Maggs AM, Bennett PM, Pinder JC, Phillips GW. The
CKK domain (DUF1781) binds microtubules and defines the CAMSAP/ssp4 family of
animal proteins. Mol Biol Evol 26: 2005–2014, 2009.
23. Baird DH, Myers KA, Mogensen M, Moss D, Baas PW. Distribution of the microtubule-related protein ninein in developing neurons. Neuropharmacology 47: 677– 683,
2004.
42. Caceres A, Kosik KS. Inhibition of neurite polarity by tau antisense oligonucleotides in
primary cerebellar neurons. Nature 343: 461– 463, 1990.
43. Caceres A, Mautino J, Kosik KS. Suppression of MAP2 in cultured cerebellar macroneurons inhibits minor neurite formation. Neuron 9: 607– 618, 1992.
44. Cappello S, Monzo P, Vallee RB. NudC is required for interkinetic nuclear migration
and neuronal migration during neocortical development. Dev Biol 357: 326 –335,
2011.
45. Chang DT, Honick AS, Reynolds IJ. Mitochondrial trafficking to synapses in cultured
primary cortical neurons. J Neurosci 26: 7035–7045, 2006.
24. Ballif BA, Arnaud L, Arthur WT, Guris D, Imamoto A, Cooper JA. Activation of a
Dab1/CrkL/C3G/Rap1 pathway in Reelin-stimulated neurons. Curr Biol 14: 606 – 610,
2004.
46. Chardin P. Function and regulation of Rnd proteins. Nat Rev Mol Cell Biol 7: 54 – 62,
2006.
25. Bamburg JR, Harris HE, Weeds AG. Partial purification and characterization of an actin
depolymerizing factor from brain. FEBS Lett 121: 178 –182, 1980.
47. Chen S, Chen J, Shi H, Wei M, Castaneda-Castellanos DR, Bultje RS, Pei X, Kriegstein
AR, Zhang M, Shi SH. Regulation of microtubule stability and organization by mammalian Par3 in specifying neuronal polarity. Dev Cell 24: 26 – 40, 2013.
27. Bartolini F, Gundersen GG. Generation of noncentrosomal microtubule arrays. J Cell
Sci 119: 4155– 4163, 2006.
28. Baudet ML, Zivraj KH, Abreu-Goodger C, Muldal A, Armisen J, Blenkiron C, Goldstein LD, Miska EA, Holt CE. miR-124 acts through CoREST to control onset of
Sema3A sensitivity in navigating retinal growth cones. Nat Neurosci 15: 29 –38, 2012.
29. Bellion A, Baudoin JP, Alvarez C, Bornens M, Metin C. Nucleokinesis in tangentially
migrating neurons comprises two alternating phases: forward migration of the Golgi/
centrosome associated with centrosome splitting and myosin contraction at the rear.
J Neurosci 25: 5691–5699, 2005.
48. Chen YM, Wang QJ, Hu HS, Yu PC, Zhu J, Drewes G, Piwnica-Worms H, Luo ZG.
Microtubule affinity-regulating kinase 2 functions downstream of the PAR-3/PAR-6/
atypical PKC complex in regulating hippocampal neuronal polarity. Proc Natl Acad Sci
USA 103: 8534 – 8539, 2006.
49. Cheng PL, Lu H, Shelly M, Gao H, Poo MM. Phosphorylation of E3 ligase Smurf1
switches its substrate preference in support of axon development. Neuron 69: 231–
243, 2011.
50. Chubb JE, Bradshaw NJ, Soares DC, Porteous DJ, Millar JK. The DISC locus in psychiatric illness. Mol Psychiatry 13: 36 – 64, 2008.
51. Conde C, Arias C, Robin M, Li A, Saito M, Chuang JZ, Nairn AC, Sung CH, Caceres A.
Evidence for the involvement of Lfc and Tctex-1 in axon formation. J Neurosci 30:
6793– 6800, 2010.
30. Bennett V, Healy J. Organizing the fluid membrane bilayer: diseases linked to spectrin
and ankyrin. Trends Mol Med 14: 28 –36, 2008.
52. Craig AM, Banker G. Neuronal polarity. Annu Rev Neurosci 17: 267–310, 1994.
31. Biernat J, Gustke N, Drewes G, Mandelkow EM, Mandelkow E. Phosphorylation of
Ser262 strongly reduces binding of tau to microtubules: distinction between PHF-like
immunoreactivity and microtubule binding. Neuron 11: 153–163, 1993.
53. Da Silva JS, Medina M, Zuliani C, Di Nardo A, Witke W, Dotti CG. RhoA/ROCK
regulation of neuritogenesis via profilin IIa-mediated control of actin stability. J Cell Biol
162: 1267–1279, 2003.
32. Bloom GS, Wagner MC, Pfister KK, Brady ST. Native structure and physical properties of bovine brain kinesin and identification of the ATP-binding subunit polypeptide.
Biochemistry 27: 3409 –3416, 1988.
54. Dajas-Bailador F, Jones EV, Whitmarsh AJ. The JIP1 scaffold protein regulates axonal
development in cortical neurons. Curr Biol 18: 221–226, 2008.
33. Borrell V, Reillo I. Emerging roles of neural stem cells in cerebral cortex development
and evolution. Dev Neurobiol 72: 955–971, 2012.
55. Davare MA, Fortin DA, Saneyoshi T, Nygaard S, Kaech S, Banker G, Soderling TR,
Wayman GA. Transient receptor potential canonical 5 channels activate Ca2⫹/calmodulin kinase Igamma to promote axon formation in hippocampal neurons. J Neurosci 29: 9794 –9808, 2009.
34. Bouckson-Castaing V, Moudjou M, Ferguson DJ, Mucklow S, Belkaid Y, Milon G,
Crocker PR. Molecular characterisation of ninein, a new coiled-coil protein of the
centrosome. J Cell Sci 109: 179 –190, 1996.
56. De Anda FC, Pollarolo G, Da Silva JS, Camoletto PG, Feiguin F, Dotti CG. Centrosome localization determines neuronal polarity. Nature 436: 704 –708, 2005.
35. Bowman AB, Kamal A, Ritchings BW, Philp AV, McGrail M, Gindhart JG, Goldstein LS.
Kinesin-dependent axonal transport is mediated by the sunday driver (SYD) protein.
Cell 103: 583–594, 2000.
57. De la Torre-Ubieta L, Gaudilliere B, Yang Y, Ikeuchi Y, Yamada T, DiBacco S, Stegmuller J, Schuller U, Salih DA, Rowitch D, Brunet A, Bonni A. A FOXO-Pak1 transcriptional pathway controls neuronal polarity. Genes Dev 24: 799 – 813, 2010.
36. Boycott KM, Flavelle S, Bureau A, Glass HC, Fujiwara TM, Wirrell E, Davey K, Chudley AE, Scott JN, McLeod DR, Parboosingh JS. Homozygous deletion of the very low
density lipoprotein receptor gene causes autosomal recessive cerebellar hypoplasia
with cerebral gyral simplification. Am J Hum Genet 77: 477– 483, 2005.
58. Delaney MK, Liu J, Zheng Y, Berndt MC, Du X. The role of Rac1 in glycoprotein
Ib-IX-mediated signal transduction and integrin activation. Arterioscler Thromb Vasc
Biol 32: 2761–2768, 2012.
37. Bradke F, Dotti CG. The role of local actin instability in axon formation. Science 283:
1931–1934, 1999.
38. Brickley K, Smith MJ, Beck M, Stephenson FA. GRIF-1 and OIP106, members of a
novel gene family of coiled-coil domain proteins: association in vivo and in vitro with
kinesin. J Biol Chem 280: 14723–14732, 2005.
39. Burton PR, Paige JL. Polarity of axoplasmic microtubules in the olfactory nerve of the
frog. Proc Natl Acad Sci USA 78: 3269 –3273, 1981.
59. Dent EW, Gupton SL, Gertler FB. The growth cone cytoskeleton in axon outgrowth
and guidance. Cold Spring Harb Perspect Biol 3: 2011.
60. Des Portes V, Pinard JM, Billuart P, Vinet MC, Koulakoff A, Carrie A, Gelot A, Dupuis
E, Motte J, Berwald-Netter Y, Catala M, Kahn A, Beldjord C, Chelly J. A novel CNS
gene required for neuronal migration and involved in X-linked subcortical laminar
heterotopia and lissencephaly syndrome. Cell 92: 51– 61, 1998.
61. Dickens M, Rogers JS, Cavanagh J, Raitano A, Xia Z, Halpern JR, Greenberg ME,
Sawyers CL, Davis RJ. A cytoplasmic inhibitor of the JNK signal transduction pathway.
Science 277: 693– 696, 1997.
40. Byrd DT, Kawasaki M, Walcoff M, Hisamoto N, Matsumoto K, Jin Y. UNC-16, a
JNK-signaling scaffold protein, regulates vesicle transport in C. elegans. Neuron 32:
787– 800, 2001.
62. Dickson B, Sprenger F, Morrison D, Hafen E. Raf functions downstream of Ras1 in the
Sevenless signal transduction pathway. Nature 360: 600 – 603, 1992.
41. Caceres A, Banker GA, Binder L. Immunocytochemical localization of tubulin and
microtubule-associated protein 2 during the development of hippocampal neurons in
culture. J Neurosci 6: 714 –722, 1986.
63. Dietrich KA, Sindelar CV, Brewer PD, Downing KH, Cremo CR, Rice SE. The kinesin-1 motor protein is regulated by a direct interaction of its head and tail. Proc Natl
Acad Sci USA 105: 8938 – 8943, 2008.
1016
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
26. Barkovich AJ, Guerrini R, Kuzniecky RI, Jackson GD, Dobyns WB. A developmental
and genetic classification for malformations of cortical development: update 2012.
Brain 135: 1348 –1369, 2012.
NAMBA ET AL.
64. Dobyns WB, Curry CJ, Hoyme HE, Turlington L, Ledbetter DH. Clinical and molecular diagnosis of Miller-Dieker syndrome. Am J Hum Genet 48: 584 –594, 1991.
65. Dotti CG, Banker GA. Experimentally induced alteration in the polarity of developing
neurons. Nature 330: 254 –256, 1987.
66. Dotti CG, Sullivan CA, Banker GA. The establishment of polarity by hippocampal
neurons in culture. J Neurosci 8: 1454 –1468, 1988.
67. Dow LE, Humbert PO. Polarity regulators and the control of epithelial architecture,
cell migration, and tumorigenesis. Int Rev Cytol 262: 253–302, 2007.
68. Drechsel DN, Hyman AA, Cobb MH, Kirschner MW. Modulation of the dynamic
instability of tubulin assembly by the microtubule-associated protein tau. Mol Biol Cell
3: 1141–1154, 1992.
69. Drewes G, Ebneth A, Preuss U, Mandelkow EM, Mandelkow E. MARK, a novel family
of protein kinases that phosphorylate microtubule-associated proteins and trigger
microtubule disruption. Cell 89: 297–308, 1997.
71. Erck C, Peris L, Andrieux A, Meissirel C, Gruber AD, Vernet M, Schweitzer A, Saoudi
Y, Pointu H, Bosc C, Salin PA, Job D, Wehland J. A vital role of tubulin-tyrosine-ligase
for neuronal organization. Proc Natl Acad Sci USA 102: 7853–7858, 2005.
72. Etemad-Moghadam B, Guo S, Kemphues KJ. Asymmetrically distributed PAR-3 protein contributes to cell polarity and spindle alignment in early C. elegans embryos. Cell
83: 743–752, 1995.
73. Etienne-Manneville S. Microtubules in cell migration. Annu Rev Cell Dev Biol 29: 471–
499, 2013.
74. Ferreira A, Busciglio J, Caceres A. Microtubule formation and neurite growth in
cerebellar macroneurons which develop in vitro: evidence for the involvement of the
microtubule-associated proteins, MAP-1a, HMW-MAP2 and Tau. Brain Res 49: 215–
228, 1989.
75. Fivaz M, Bandara S, Inoue T, Meyer T. Robust neuronal symmetry breaking by Rastriggered local positive feedback. Curr Biol 18: 44 –50, 2008.
76. Florio M, Huttner WB. Neural progenitors, neurogenesis and the evolution of the
neocortex. Development 141: 2182–2194, 2014.
77. Flynn KC, Hellal F, Neukirchen D, Jacob S, Tahirovic S, Dupraz S, Stern S, Garvalov
BK, Gurniak C, Shaw AE, Meyn L, Wedlich-Soldner R, Bamburg JR, Small JV, Witke W,
Bradke F. ADF/cofilin-mediated actin retrograde flow directs neurite formation in the
developing brain. Neuron 76: 1091–1107, 2012.
78. Foster R, Hu KQ, Lu Y, Nolan KM, Thissen J, Settleman J. Identification of a novel
human Rho protein with unusual properties: GTPase deficiency and in vivo farnesylation. Mol Cell Biol 16: 2689 –2699, 1996.
79. Fourniol FJ, Sindelar CV, Amigues B, Clare DK, Thomas G, Perderiset M, Francis F,
Houdusse A, Moores CA. Template-free 13-protofilament microtubule-MAP assembly visualized at 8 A resolution. J Cell Biol 191: 463– 470, 2010.
80. Francis F, Koulakoff A, Boucher D, Chafey P, Schaar B, Vinet MC, Friocourt G,
McDonnell N, Reiner O, Kahn A, McConnell SK, Berwald-Netter Y, Denoulet P,
Chelly J. Doublecortin is a developmentally regulated, microtubule-associated protein
expressed in migrating and differentiating neurons. Neuron 23: 247–256, 1999.
81. Franco SJ, Martinez-Garay I, Gil-Sanz C, Harkins-Perry SR, Muller U. Reelin regulates
cadherin function via Dab1/Rap1 to control neuronal migration and lamination in the
neocortex. Neuron 69: 482– 497, 2011.
82. Fransson S, Ruusala A, Aspenstrom P. The atypical Rho GTPases Miro-1 and Miro-2
have essential roles in mitochondrial trafficking. Biochem Biophys Res Commun 344:
500 –510, 2006.
83. Friedl P, Alexander S. Cancer invasion and the microenvironment: plasticity and
reciprocity. Cell 147: 992–1009, 2011.
84. Fuentes P, Canovas J, Berndt FA, Noctor SC, Kukuljan M. CoREST/LSD1 control the
development of pyramidal cortical neurons. Cereb Cortex 22: 1431–1441, 2012.
86. Fukuda M. Rab27 effectors, pleiotropic regulators in secretory pathways. Traffic 14:
949 –963, 2013.
87. Fukuda M. Regulation of secretory vesicle traffic by Rab small GTPases. Cell Mol Life
Sci 65: 2801–2813, 2008.
88. Funahashi Y, Namba T, Fujisue S, Itoh N, Nakamuta S, Kato K, Shimada A, Xu C, Shan
W, Nishioka T, Kaibuchi K. ERK2-mediated phosphorylation of Par3 regulates neuronal polarization. J Neurosci 33: 13270 –13285, 2013.
89. Galiano MR, Jha S, Ho TS, Zhang C, Ogawa Y, Chang KJ, Stankewich MC, Mohler PJ,
Rasband MN. A distal axonal cytoskeleton forms an intra-axonal boundary that controls axon initial segment assembly. Cell 149: 1125–1139, 2012.
90. Garnham CP, Roll-Mecak A. The chemical complexity of cellular microtubules: tubulin post-translational modification enzymes and their roles in tuning microtubule functions. Cytoskeleton 69: 442– 463, 2012.
91. Gartner A, Fornasiero EF, Dotti CG. N-cadherin: a new player in neuronal polarity.
Cell Cycle 11: 2223–2224, 2012.
92. Gartner A, Fornasiero EF, Munck S, Vennekens K, Seuntjens E, Huttner WB, Valtorta
F, Dotti CG. N-cadherin specifies first asymmetry in developing neurons. EMBO J 31:
1893–1903, 2012.
93. Garvalov BK, Flynn KC, Neukirchen D, Meyn L, Teusch N, Wu X, Brakebusch C,
Bamburg JR, Bradke F. Cdc42 regulates cofilin during the establishment of neuronal
polarity. J Neurosci 27: 13117–13129, 2007.
94. Gaughwin P, Ciesla M, Yang H, Lim B, Brundin P. Stage-specific modulation of cortical
neuronal development by Mmu-miR-134. Cereb Cortex 21: 1857–1869, 2011.
95. Ge W, He F, Kim KJ, Blanchi B, Coskun V, Nguyen L, Wu X, Zhao J, Heng JI,
Martinowich K, Tao J, Wu H, Castro D, Sobeih MM, Corfas G, Gleeson JG, Greenberg
ME, Guillemot F, Sun YE. Coupling of cell migration with neurogenesis by proneural
bHLH factors. Proc Natl Acad Sci USA 103: 1319 –1324, 2006.
96. Gertler FB, Doctor JS, Hoffmann FM. Genetic suppression of mutations in the Drosophila abl proto-oncogene homolog. Science 248: 857– 860, 1990.
97. Glater EE, Megeath LJ, Stowers RS, Schwarz TL. Axonal transport of mitochondria
requires milton to recruit kinesin heavy chain and is light chain independent. J Cell Biol
173: 545–557, 2006.
98. Gleeson JG, Allen KM, Fox JW, Lamperti ED, Berkovic S, Scheffer I, Cooper EC,
Dobyns WB, Minnerath SR, Ross ME, Walsh CA. Doublecortin, a brain-specific gene
mutated in human X-linked lissencephaly and double cortex syndrome, encodes a
putative signaling protein. Cell 92: 63–72, 1998.
99. Goldberg DJ, Burmeister DW. Stages in axon formation: observations of growth of
Aplysia axons in culture using video-enhanced contrast-differential interference contrast microscopy. J Cell Biol 103: 1921–1931, 1986.
100. Goldstein B, Macara IG. The PAR proteins: fundamental players in animal cell polarization. Dev Cell 13: 609 – 622, 2007.
101. Gomis-Ruth S, Wierenga CJ, Bradke F. Plasticity of polarization: changing dendrites
into axons in neurons integrated in neuronal circuits. Curr Biol 18: 992–1000, 2008.
102. Goshima Y, Nakamura F, Strittmatter P, Strittmatter SM. Collapsin-induced growth
cone collapse mediated by an intracellular protein related to UNC-33. Nature 376:
509 –514, 1995.
103. Goslin K, Banker G. Experimental observations on the development of polarity by
hippocampal neurons in culture. J Cell Biol 108: 1507–1516, 1989.
104. Graham ME, Ruma-Haynes P, Capes-Davis AG, Dunn JM, Tan TC, Valova VA, Robinson PJ, Jeffrey PL. Multisite phosphorylation of doublecortin by cyclin-dependent
kinase 5. Biochem J 381: 471– 481, 2004.
105. Guasch RM, Scambler P, Jones GE, Ridley AJ. RhoE regulates actin cytoskeleton
organization and cell migration. Mol Cell Biol 18: 4761– 4771, 1998.
106. Gubba EM, Fawcett JW, Herbert J. The effects of corticosterone and dehydroepiandrosterone on neurotrophic factor mRNA expression in primary hippocampal and
astrocyte cultures. Brain Res 127: 48 –59, 2004.
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
1017
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
70. Enomoto A, Asai N, Namba T, Wang Y, Kato T, Tanaka M, Tatsumi H, Taya S, Tsuboi
D, Kuroda K, Kaneko N, Sawamoto K, Miyamoto R, Jijiwa M, Murakumo Y, Sokabe M,
Seki T, Kaibuchi K, Takahashi M. Roles of disrupted-in-schizophrenia 1-interacting
protein girdin in postnatal development of the dentate gyrus. Neuron 63: 774 –787,
2009.
85. Fukata Y, Itoh TJ, Kimura T, Menager C, Nishimura T, Shiromizu T, Watanabe H,
Inagaki N, Iwamatsu A, Hotani H, Kaibuchi K. CRMP-2 binds to tubulin heterodimers
to promote microtubule assembly. Nat Cell Biol 4: 583–591, 2002.
NEURONAL POLARITY SIGNALING
107. Guillaud L, Wong R, Hirokawa N. Disruption of KIF17-Mint1 interaction by CaMKIIdependent phosphorylation: a molecular model of kinesin-cargo release. Nat Cell Biol
10: 19 –29, 2008.
108. Guillemot F. Cell fate specification in the mammalian telencephalon. Prog Neurobiol
83: 37–52, 2007.
109. Guo S, Kemphues KJ. par-1, a gene required for establishing polarity in C. elegans
embryos, encodes a putative Ser/Thr kinase that is asymmetrically distributed. Cell 81:
611– 620, 1995.
110. Guo X, Macleod GT, Wellington A, Hu F, Panchumarthi S, Schoenfield M, Marin L,
Charlton MP, Atwood HL, Zinsmaier KE. The GTPase dMiro is required for axonal
transport of mitochondria to Drosophila synapses. Neuron 47: 379 –393, 2005.
111. Gupton SL, Gertler FB. Integrin signaling switches the cytoskeletal and exocytic machinery that drives neuritogenesis. Dev Cell 18: 725–736, 2010.
112. Haffner C, Jarchau T, Reinhard M, Hoppe J, Lohmann SM, Walter U. Molecular
cloning, structural analysis and functional expression of the proline-rich focal adhesion
and microfilament-associated protein VASP. EMBO J 14: 19 –27, 1995.
114. Hall DH, Hedgecock EM. Kinesin-related gene unc-104 is required for axonal transport of synaptic vesicles in C. elegans. Cell 65: 837– 847, 1991.
115. Hamelers IH, Olivo C, Mertens AE, Pegtel DM, van der Kammen RA, Sonnenberg A,
Collard JG. The Rac activator Tiam1 is required for (alpha)3(beta)1-mediated
laminin-5 deposition, cell spreading, and cell migration. J Cell Biol 171: 871– 881, 2005.
116. Hammond JW, Huang CF, Kaech S, Jacobson C, Banker G, Verhey KJ. Posttranslational modifications of tubulin and the polarized transport of kinesin-1 in neurons. Mol
Biol Cell 21: 572–583, 2010.
117. Hand R, Bortone D, Mattar P, Nguyen L, Heng JI, Guerrier S, Boutt E, Peters E, Barnes
AP, Parras C, Schuurmans C, Guillemot F, Polleux F. Phosphorylation of Neurogenin2
specifies the migration properties and the dendritic morphology of pyramidal neurons
in the neocortex. Neuron 48: 45– 62, 2005.
118. Hartman JJ, Mahr J, McNally K, Okawa K, Iwamatsu A, Thomas S, Cheesman S,
Heuser J, Vale RD, McNally FJ. Katanin, a microtubule-severing protein, is a novel
AAA ATPase that targets to the centrosome using a WD40-containing subunit. Cell
93: 277–287, 1998.
119. Hatanaka Y, Yamauchi K. Excitatory cortical neurons with multipolar shape establish
neuronal polarity by forming a tangentially oriented axon in the intermediate zone.
Cereb Cortex 23: 105–113, 2013.
120. Hatta K, Takeichi M. Expression of N-cadherin adhesion molecules associated with
early morphogenetic events in chick development. Nature 320: 447– 449, 1986.
121. Hedgecock EM, Culotti JG, Thomson JN, Perkins LA. Axonal guidance mutants of
Caenorhabditis elegans identified by filling sensory neurons with fluorescein dyes. Dev
Biol 111: 158 –170, 1985.
122. Hedstrom KL, Ogawa Y, Rasband MN. AnkyrinG is required for maintenance of the
axon initial segment and neuronal polarity. J Cell Biol 183: 635– 640, 2008.
123. Heidemann SR, Landers JM, Hamborg MA. Polarity orientation of axonal microtubules. J Cell Biol 91: 661– 665, 1981.
124. Hellal F, Hurtado A, Ruschel J, Flynn KC, Laskowski CJ, Umlauf M, Kapitein LC, Strikis
D, Lemmon V, Bixby J, Hoogenraad CC, Bradke F. Microtubule stabilization reduces
scarring and causes axon regeneration after spinal cord injury. Science 331: 928 –931,
2011.
125. Heng JI, Nguyen L, Castro DS, Zimmer C, Wildner H, Armant O, SkowronskaKrawczyk D, Bedogni F, Matter JM, Hevner R, Guillemot F. Neurogenin 2 controls
cortical neuron migration through regulation of Rnd2. Nature 455: 114 –118, 2008.
129. Hirai S, Banba Y, Satake T, Ohno S. Axon formation in neocortical neurons depends
on stage-specific regulation of microtubule stability by the dual leucine zipper kinasec-Jun N-terminal kinase pathway. J Neurosci 31: 6468 – 6480, 2011.
130. Hirano S, Takeichi M. Cadherins in brain morphogenesis and wiring. Physiol Rev 92:
597– 634, 2012.
131. Hirokawa N, Noda Y, Tanaka Y, Niwa S. Kinesin superfamily motor proteins and
intracellular transport. Nat Rev Mol Cell Biol 10: 682– 696, 2009.
132. Hirokawa N, Takemura R. Molecular motors and mechanisms of directional transport
in neurons. Nat Rev Neurosci 6: 201–214, 2005.
133. Hirotsune S, Fleck MW, Gambello MJ, Bix GJ, Chen A, Clark GD, Ledbetter DH,
McBain CJ, Wynshaw-Boris A. Graded reduction of Pafah1b1 (Lis1) activity results in
neuronal migration defects and early embryonic lethality. Nat Genet 19: 333–339,
1998.
134. Hogan C, Serpente N, Cogram P, Hosking CR, Bialucha CU, Feller SM, Braga VM,
Birchmeier W, Fujita Y. Rap1 regulates the formation of E-cadherin-based cell-cell
contacts. Mol Cell Biol 24: 6690 – 6700, 2004.
135. Hollenbeck PJ, Saxton WM. The axonal transport of mitochondria. J Cell Sci 118:
5411–5419, 2005.
136. Homma N, Takei Y, Tanaka Y, Nakata T, Terada S, Kikkawa M, Noda Y, Hirokawa N.
Kinesin superfamily protein 2A (KIF2A) functions in suppression of collateral branch
extension. Cell 114: 229 –239, 2003.
137. Hong SE, Shugart YY, Huang DT, Shahwan SA, Grant PE, Hourihane JO, Martin ND,
Walsh CA. Autosomal recessive lissencephaly with cerebellar hypoplasia is associated
with human RELN mutations. Nat Genet 26: 93–96, 2000.
138. Hoogenraad CC, Feliu-Mojer MI, Spangler SA, Milstein AD, Dunah AW, Hung AY,
Sheng M. Liprinalpha1 degradation by calcium/calmodulin-dependent protein kinase II
regulates LAR receptor tyrosine phosphatase distribution and dendrite development.
Dev Cell 12: 587– 602, 2007.
139. Horiguchi K, Hanada T, Fukui Y, Chishti AH. Transport of PIP3 by GAKIN, a kinesin-3
family protein, regulates neuronal cell polarity. J Cell Biol 174: 425– 436, 2006.
140. Horiuchi D, Barkus RV, Pilling AD, Gassman A, Saxton WM. APLIP1, a kinesin binding
JIP-1/JNK scaffold protein, influences the axonal transport of both vesicles and mitochondria in Drosophila. Curr Biol 15: 2137–2141, 2005.
141. Horiuchi D, Collins CA, Bhat P, Barkus RV, Diantonio A, Saxton WM. Control of a
kinesin-cargo linkage mechanism by JNK pathway kinases. Curr Biol 17: 1313–1317,
2007.
142. Hoshino T, Sakisaka T, Baba T, Yamada T, Kimura T, Takai Y. Regulation of Ecadherin endocytosis by nectin through afadin, Rap1, and p120ctn. J Biol Chem 280:
24095–24103, 2005.
143. Hsu YC, Fuchs E. A family business: stem cell progeny join the niche to regulate
homeostasis. Nat Rev Mol Cell Biol 13: 103–114, 2012.
144. Huang CF, Banker G. The translocation selectivity of the kinesins that mediate neuronal organelle transport. Traffic 13: 549 –564, 2012.
145. Huang SH, Duan S, Sun T, Wang J, Zhao L, Geng Z, Yan J, Sun HJ, Chen ZY. JIP3
mediates TrkB axonal anterograde transport and enhances BDNF signaling by directly
bridging TrkB with kinesin-1. J Neurosci 31: 10602–10614, 2011.
146. Hubbert C, Guardiola A, Shao R, Kawaguchi Y, Ito A, Nixon A, Yoshida M, Wang XF,
Yao TP. HDAC6 is a microtubule-associated deacetylase. Nature 417: 455– 458,
2002.
147. Hudish LI, Blasky AJ, Appel B. miR-219 regulates neural precursor differentiation by
direct inhibition of apical par polarity proteins. Dev Cell 27: 387–398, 2013.
126. Heng JI, Qu Z, Ohtaka-Maruyama C, Okado H, Kasai M, Castro D, Guillemot F, Tan
SS. The zinc finger transcription factor RP58 negatively regulates Rnd2 for the control
of neuronal migration during cerebral cortical development. Cereb Cortex 25: 806 –
816, 2015.
148. Inagaki N, Chihara K, Arimura N, Menager C, Kawano Y, Matsuo N, Nishimura T,
Amano M, Kaibuchi K. CRMP-2 induces axons in cultured hippocampal neurons. Nat
Neurosci 4: 781–782, 2001.
127. Hilliard MA, Bargmann CI. Wnt signals and frizzled activity orient anterior-posterior
axon outgrowth in C. elegans. Dev Cell 10: 379 –390, 2006.
149. Inagaki N, Toriyama M, Sakumura Y. Systems biology of symmetry breaking during
neuronal polarity formation. Dev Neurobiol 71: 584 –593, 2011.
1018
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
113. Hall A, Paterson HF, Adamson P, Ridley AJ. Cellular responses regulated by rhorelated small GTP-binding proteins. Philos Trans R Soc Lond B Biol Sci 340: 267–271,
1993.
128. Hirabayashi Y, Gotoh Y. Epigenetic control of neural precursor cell fate during development. Nat Rev Neurosci 11: 377–388, 2010.
NAMBA ET AL.
150. Inomata H, Nakamura Y, Hayakawa A, Takata H, Suzuki T, Miyazawa K, Kitamura N.
A scaffold protein JIP-1b enhances amyloid precursor protein phosphorylation by JNK
and its association with kinesin light chain 1. J Biol Chem 278: 22946 –22955, 2003.
151. Ip JP, Shi L, Chen Y, Itoh Y, Fu WY, Betz A, Yung WH, Gotoh Y, Fu AK, Ip NY.
Alpha2-chimaerin controls neuronal migration and functioning of the cerebral cortex
through CRMP-2. Nat Neurosci 15: 39 – 47, 2012.
152. Izumi Y, Hirose T, Tamai Y, Hirai S, Nagashima Y, Fujimoto T, Tabuse Y, Kemphues
KJ, Ohno S. An atypical PKC directly associates and colocalizes at the epithelial tight
junction with ASIP, a mammalian homologue of Caenorhabditis elegans polarity protein
PAR-3. J Cell Biol 143: 95–106, 1998.
153. Jaaro-Peled H, Hayashi-Takagi A, Seshadri S, Kamiya A, Brandon NJ, Sawa A. Neurodevelopmental mechanisms of schizophrenia: understanding disturbed postnatal
brain maturation through neuregulin-1-ErbB4 and DISC1. Trends Neurosci 32: 485–
495, 2009.
154. Jiang H, Guo W, Liang X, Rao Y. Both the establishment and the maintenance of
neuronal polarity require active mechanisms: critical roles of GSK-3beta and its upstream regulators. Cell 120: 123–135, 2005.
156. Joberty G, Petersen C, Gao L, Macara IG. The cell-polarity protein Par6 links Par3 and
atypical protein kinase C to Cdc42. Nat Cell Biol 2: 531–539, 2000.
157. Johansson A, Driessens M, Aspenstrom P. The mammalian homologue of the Caenorhabditis elegans polarity protein PAR-6 is a binding partner for the Rho GTPases
Cdc42 and Rac1. J Cell Sci 113: 3267–3275, 2000.
158. Jossin Y, Cooper JA. Reelin, Rap1 and N-cadherin orient the migration of multipolar
neurons in the developing neocortex. Nat Neurosci 14: 697–703, 2011.
159. Jossin Y, Gui L, Goffinet AM. Processing of Reelin by embryonic neurons is important
for function in tissue but not in dissociated cultured neurons. J Neurosci 27: 4243–
4252, 2007.
160. Kaech S, Ludin B, Matus A. Cytoskeletal plasticity in cells expressing neuronal microtubule-associated proteins. Neuron 17: 1189 –1199, 1996.
161. Kalebic N, Sorrentino S, Perlas E, Bolasco G, Martinez C, Heppenstall PA. alphaTAT1
is the major alpha-tubulin acetyltransferase in mice. Nature Commun 4: 1962, 2013.
162. Kamiya A, Kubo K, Tomoda T, Takaki M, Youn R, Ozeki Y, Sawamura N, Park U,
Kudo C, Okawa M, Ross CA, Hatten ME, Nakajima K, Sawa A. A schizophreniaassociated mutation of DISC1 perturbs cerebral cortex development. Nat Cell Biol 7:
1167–1178, 2005.
163. Karagogeos D. Neural GPI-anchored cell adhesion molecules. Front Biosci 8: 1304 –
1320, 2003.
164. Kasahara K, Watanabe K, Kozutsumi Y, Oohira A, Yamamoto T, Sanai Y. Association
of GPI-anchored protein TAG-1 with src-family kinase Lyn in lipid rafts of cerebellar
granule cells. Neurochem Res 27: 823– 829, 2002.
165. Kasahara K, Watanabe K, Takeuchi K, Kaneko H, Oohira A, Yamamoto T, Sanai Y.
Involvement of gangliosides in glycosylphosphatidylinositol-anchored neuronal cell
adhesion molecule TAG-1 signaling in lipid rafts. J Biol Chem 275: 34701–34709, 2000.
166. Kawano Y, Yoshimura T, Tsuboi D, Kawabata S, Kaneko-Kawano T, Shirataki H,
Takenawa T, Kaibuchi K. CRMP-2 is involved in kinesin-1-dependent transport of the
Sra-1/WAVE1 complex and axon formation. Mol Cell Biol 25: 9920 –9935, 2005.
167. Kawauchi T, Chihama K, Nabeshima Y, Hoshino M. The in vivo roles of STEF/Tiam1,
Rac1 and JNK in cortical neuronal migration. EMBO J 22: 4190 – 4201, 2003.
168. Kawauchi T, Sekine K, Shikanai M, Chihama K, Tomita K, Kubo K, Nakajima K,
Nabeshima Y, Hoshino M. Rab GTPases-dependent endocytic pathways regulate
neuronal migration and maturation through N-cadherin trafficking. Neuron 67: 588 –
602, 2010.
169. Keays DA, Tian G, Poirier K, Huang GJ, Siebold C, Cleak J, Oliver PL, Fray M, Harvey
RJ, Molnar Z, Pinon MC, Dear N, Valdar W, Brown SD, Davies KE, Rawlins JN, Cowan
NJ, Nolan P, Chelly J, Flint J. Mutations in alpha-tubulin cause abnormal neuronal
migration in mice and lissencephaly in humans. Cell 128: 45–57, 2007.
171. Kennedy NJ, Martin G, Ehrhardt AG, Cavanagh-Kyros J, Kuan CY, Rakic P, Flavell RA,
Treistman SN, Davis RJ. Requirement of JIP scaffold proteins for NMDA-mediated
signal transduction. Genes Dev 21: 2336 –2346, 2007.
172. Kim JY, Duan X, Liu CY, Jang MH, Guo JU, Pow-anpongkul N, Kang E, Song H, Ming
GL. DISC1 regulates new neuron development in the adult brain via modulation of
AKT-mTOR signaling through KIAA1212. Neuron 63: 761–773, 2009.
173. Kimura K, Ito M, Amano M, Chihara K, Fukata Y, Nakafuku M, Yamamori B, Feng J,
Nakano T, Okawa K, Iwamatsu A, Kaibuchi K. Regulation of myosin phosphatase by
Rho and Rho-associated kinase (Rho-kinase). Science 273: 245–248, 1996.
174. Kimura T, Arimura N, Fukata Y, Watanabe H, Iwamatsu A, Kaibuchi K. Tubulin and
CRMP-2 complex is transported via Kinesin-1. J Neurochem 93: 1371–1382, 2005.
175. Kishimoto T, Itoh K, Umekage M, Tonosaki M, Yaoi T, Fukui K, Lemmon VP, Fushiki
S. Downregulation of L1 perturbs neuronal migration and alters the expression of
transcription factors in murine neocortex. J Neurosci Res 91: 42–50, 2013.
176. Knoblich JA. Asymmetric cell division during animal development. Nat Rev Mol Cell Biol
2: 11–20, 2001.
177. Koehnle TJ, Brown A. Slow axonal transport of neurofilament protein in cultured
neurons. J Cell Biol 144: 447– 458, 1999.
178. Konishi Y, Setou M. Tubulin tyrosination navigates the kinesin-1 motor domain to
axons. Nat Neurosci 12: 559 –567, 2009.
179. Konno D, Yoshimura S, Hori K, Maruoka H, Sobue K. Involvement of the phosphatidylinositol 3-kinase/rac1 and cdc42 pathways in radial migration of cortical neurons. J
Biol Chem 280: 5082–5088, 2005.
180. Kosako H, Gotoh Y, Matsuda S, Ishikawa M, Nishida E. Xenopus MAP kinase activator
is a serine/threonine/tyrosine kinase activated by threonine phosphorylation. EMBO J
11: 2903–2908, 1992.
181. Koushika SP. “JIP”ing along the axon: the complex roles of JIPs in axonal transport.
Bioessays 30: 10 –14, 2008.
182. Kumar RA, Pilz DT, Babatz TD, Cushion TD, Harvey K, Topf M, Yates L, Robb S,
Uyanik G, Mancini GM, Rees MI, Harvey RJ, Dobyns WB. TUBA1A mutations cause
wide spectrum lissencephaly (smooth brain) and suggest that multiple neuronal migration pathways converge on alpha tubulins. Hum Mol Genet 19: 2817–2827, 2010.
183. Kuznetsov SA, Vaisberg EA, Shanina NA, Magretova NN, Chernyak VY, Gelfand VI.
The quaternary structure of bovine brain kinesin. EMBO J 7: 353–356, 1988.
184. Kwiatkowski AV, Rubinson DA, Dent EW, Edward van Veen J, Leslie JD, Zhang J,
Mebane LM, Philippar U, Pinheiro EM, Burds AA, Bronson RT, Mori S, Fassler R,
Gertler FB. Ena/VASP is required for neuritogenesis in the developing cortex. Neuron
56: 441– 455, 2007.
185. Kyriakis JM, App H, Zhang XF, Banerjee P, Brautigan DL, Rapp UR, Avruch J. Raf-1
activates MAP kinase-kinase. Nature 358: 417– 421, 1992.
186. Kyriakis JM, Banerjee P, Nikolakaki E, Dai T, Rubie EA, Ahmad MF, Avruch J,
Woodgett JR. The stress-activated protein kinase subfamily of c-Jun kinases. Nature
369: 156 –160, 1994.
187. Lamoureux P, Ruthel G, Buxbaum RE, Heidemann SR. Mechanical tension can specify
axonal fate in hippocampal neurons. J Cell Biol 159: 499 –508, 2002.
188. Ledbetter SA, Kuwano A, Dobyns WB, Ledbetter DH. Microdeletions of chromosome 17p13 as a cause of isolated lissencephaly. Am J Hum Genet 50: 182–189, 1992.
189. Leemhuis J, Bouche E, Frotscher M, Henle F, Hein L, Herz J, Meyer DK, Pichler M,
Roth G, Schwan C, Bock HH. Reelin signals through apolipoprotein E receptor 2 and
Cdc42 to increase growth cone motility and filopodia formation. J Neurosci 30:
14759 –14772, 2010.
190. Lefcort F, Bentley D. Organization of cytoskeletal elements and organelles preceding
growth cone emergence from an identified neuron in situ. J Cell Biol 108: 1737–1749,
1989.
191. Leone DP, Srinivasan K, Brakebusch C, McConnell SK. The rho GTPase Rac1 is
required for proliferation and survival of progenitors in the developing forebrain. Dev
Neurobiol 70: 659 – 678, 2010.
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
1019
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
155. Jiang K, Hua S, Mohan R, Grigoriev I, Yau KW, Liu Q, Katrukha EA, Altelaar AF, Heck
AJ, Hoogenraad CC, Akhmanova A. Microtubule minus-end stabilization by polymerization-driven CAMSAP deposition. Dev Cell 28: 295–309, 2014.
170. Kemphues KJ, Priess JR, Morton DG, Cheng NS. Identification of genes required for
cytoplasmic localization in early C. elegans embryos. Cell 52: 311–320, 1988.
NEURONAL POLARITY SIGNALING
192. Lewis TL Jr, Mao T, Svoboda K, Arnold DB. Myosin-dependent targeting of transmembrane proteins to neuronal dendrites. Nat Neurosci 12: 568 –576, 2009.
213. Miyata T, Kawaguchi A, Okano H, Ogawa M. Asymmetric inheritance of radial glial
fibers by cortical neurons. Neuron 31: 727–741, 2001.
193. Li R, Gundersen GG. Beyond polymer polarity: how the cytoskeleton builds a polarized cell. Nat Rev Mol Cell Biol 9: 860 – 873, 2008.
214. Miyata T, Kawaguchi A, Saito K, Kawano M, Muto T, Ogawa M. Asymmetric production of surface-dividing and non-surface-dividing cortical progenitor cells. Development 131: 3133–3145, 2004.
194. Li Z, He X, Feng J. Dicer is essential for neuronal polarity. Int J Dev Neurosci 30:
607– 611, 2012.
195. Lin D, Edwards AS, Fawcett JP, Mbamalu G, Scott JD, Pawson T. A mammalian
PAR-3-PAR-6 complex implicated in Cdc42/Rac1 and aPKC signalling and cell polarity.
Nat Cell Biol 2: 540 –547, 2000.
196. Lowery LA, Van Vactor D. The trip of the tip: understanding the growth cone machinery. Nat Rev Mol Cell Biol 10: 332–343, 2009.
197. Luders J, Stearns T. Microtubule-organizing centres: a re-evaluation. Nat Rev Mol Cell
Biol 8: 161–167, 2007.
198. Lui JH, Hansen DV, Kriegstein AR. Development and evolution of the human neocortex. Cell 146: 18 –36, 2011.
200. Macaskill AF, Rinholm JE, Twelvetrees AE, Arancibia-Carcamo IL, Muir J, Fransson A,
Aspenstrom P, Attwell D, Kittler JT. Miro1 is a calcium sensor for glutamate receptordependent localization of mitochondria at synapses. Neuron 61: 541–555, 2009.
201. Maekawa M, Ishizaki T, Boku S, Watanabe N, Fujita A, Iwamatsu A, Obinata T, Ohashi
K, Mizuno K, Narumiya S. Signaling from Rho to the actin cytoskeleton through
protein kinases ROCK and LIM-kinase. Science 285: 895– 898, 1999.
216. Mogensen MM, Malik A, Piel M, Bouckson-Castaing V, Bornens M. Microtubule minus-end anchorage at centrosomal and non-centrosomal sites: the role of ninein. J Cell
Sci 113: 3013–3023, 2000.
217. Mok H, Shin H, Kim S, Lee JR, Yoon J, Kim E. Association of the kinesin superfamily
motor protein KIF1Balpha with postsynaptic density-95 (PSD-95), synapse-associated protein-97, and synaptic scaffolding molecule PSD-95/discs large/zona occludens-1 proteins. J Neurosci 22: 5253–5258, 2002.
218. Moore R, Boyd L. Analysis of RING finger genes required for embryogenesis in C.
elegans. Genesis 38: 1–12, 2004.
219. Moores CA, Perderiset M, Francis F, Chelly J, Houdusse A, Milligan RA. Mechanism of
microtubule stabilization by doublecortin. Mol Cell 14: 833– 839, 2004.
220. Moreno-De-Luca A, Myers SM, Challman TD, Moreno-De-Luca D, Evans DW, Ledbetter DH. Developmental brain dysfunction: revival and expansion of old concepts
based on new genetic evidence. Lancet Neurol 12: 406 – 414, 2013.
221. Morfini G, DiTella MC, Feiguin F, Carri N, Caceres A. Neurotrophin-3 enhances
neurite outgrowth in cultured hippocampal pyramidal neurons. J Neurosci Res 39:
219 –232, 1994.
202. Mandel G, Fiondella CG, Covey MV, Lu DD, Loturco JJ, Ballas N. Repressor element
1 silencing transcription factor (REST) controls radial migration and temporal neuronal specification during neocortical development. Proc Natl Acad Sci USA 108: 16789 –
16794, 2011.
222. Morfini G, Szebenyi G, Elluru R, Ratner N, Brady ST. Glycogen synthase kinase 3
phosphorylates kinesin light chains and negatively regulates kinesin-based motility.
EMBO J 21: 281–293, 2002.
203. Manent JB, Wang Y, Chang Y, Paramasivam M, LoTurco JJ. Dcx reexpression reduces
subcortical band heterotopia and seizure threshold in an animal model of neuronal
migration disorder. Nat Med 15: 84 –90, 2009.
223. Mori D, Yamada M, Mimori-Kiyosue Y, Shirai Y, Suzuki A, Ohno S, Saya H, WynshawBoris A, Hirotsune S. An essential role of the aPKC-Aurora A-NDEL1 pathway in
neurite elongation by modulation of microtubule dynamics. Nature Cell Biol 11: 1057–
1068, 2009.
204. Mao Y, Ge X, Frank CL, Madison JM, Koehler AN, Doud MK, Tassa C, Berry EM, Soda
T, Singh KK, Biechele T, Petryshen TL, Moon RT, Haggarty SJ, Tsai LH. Disrupted in
schizophrenia 1 regulates neuronal progenitor proliferation via modulation of
GSK3beta/beta-catenin signaling. Cell 136: 1017–1031, 2009.
224. Mori K, Amano M, Takefuji M, Kato K, Morita Y, Nishioka T, Matsuura Y, Murohara T,
Kaibuchi K. Rho-kinase contributes to sustained RhoA activation through phosphorylation of p190A RhoGAP. J Biol Chem 284: 5067–5076, 2009.
205. Martin-Belmonte F, Perez-Moreno M. Epithelial cell polarity, stem cells and cancer.
Nature Rev Cancer 12: 23–38, 2012.
225. Morris SM, Albrecht U, Reiner O, Eichele G, Yu-Lee LY. The lissencephaly gene
product Lis1, a protein involved in neuronal migration, interacts with a nuclear movement protein, NudC. Curr Biol 8: 603– 606, 1998.
206. Martynoga B, Drechsel D, Guillemot F. Molecular control of neurogenesis: a view
from the mammalian cerebral cortex. Cold Spring Harb Perspect Biol 4: 2012.
207. Maussion G, Carayol J, Lepagnol-Bestel AM, Tores F, Loe-Mie Y, Milbreta U, Rousseau F, Fontaine K, Renaud J, Moalic JM, Philippi A, Chedotal A, Gorwood P, Ramoz
N, Hager J, Simonneau M. Convergent evidence identifying MAP/microtubule affinityregulating kinase 1 (MARK1) as a susceptibility gene for autism. Hum Mol Genet 17:
2541–2551, 2008.
208. McNally FJ, Okawa K, Iwamatsu A, Vale RD. Katanin, the microtubule-severing
ATPase, is concentrated at centrosomes. J Cell Sci 109: 561–567, 1996.
209. Meiri D, Greeve MA, Brunet A, Finan D, Wells CD, LaRose J, Rottapel R. Modulation
of Rho guanine exchange factor Lfc activity by protein kinase A-mediated phosphorylation. Mol Cell Biol 29: 5963–5973, 2009.
210. Menager C, Arimura N, Fukata Y, Kaibuchi K. PIP3 is involved in neuronal polarization
and axon formation. J Neurochem 89: 109 –118, 2004.
211. Mertens AE, Rygiel TP, Olivo C, van der Kammen R, Collard JG. The Rac activator
Tiam1 controls tight junction biogenesis in keratinocytes through binding to and
activation of the Par polarity complex. J Cell Biol 170: 1029 –1037, 2005.
212. Millar JK, Pickard BS, Mackie S, James R, Christie S, Buchanan SR, Malloy MP, Chubb
JE, Huston E, Baillie GS, Thomson PA, Hill EV, Brandon NJ, Rain JC, Camargo LM,
Whiting PJ, Houslay MD, Blackwood DH, Muir WJ, Porteous DJ. DISC1 and PDE4B
are interacting genetic factors in schizophrenia that regulate cAMP signaling. Science
310: 1187–1191, 2005.
1020
226. Morrow EM, Yoo SY, Flavell SW, Kim TK, Lin Y, Hill RS, Mukaddes NM, Balkhy S,
Gascon G, Hashmi A, Al-Saad S, Ware J, Joseph RM, Greenblatt R, Gleason D, Ertelt
JA, Apse KA, Bodell A, Partlow JN, Barry B, Yao H, Markianos K, Ferland RJ, Greenberg ME, Walsh CA. Identifying autism loci and genes by tracing recent shared ancestry. Science 321: 218 –223, 2008.
227. Morton DG, Roos JM, Kemphues KJ. par-4, a gene required for cytoplasmic localization and determination of specific cell types in Caenorhabditis elegans embryogenesis.
Genetics 130: 771–790, 1992.
228. Morton DG, Shakes DC, Nugent S, Dichoso D, Wang W, Golden A, Kemphues KJ.
The Caenorhabditis elegans par-5 gene encodes a 14-3-3 protein required for cellular
asymmetry in the early embryo. Dev Biol 241: 47–58, 2002.
229. Murofushi H. Purification and characterization of tubulin-tyrosine ligase from porcine
brain. J Biochem 87: 979 –984, 1980.
230. Nadarajah B. Radial glia and somal translocation of radial neurons in the developing
cerebral cortex. Glia 43: 33–36, 2003.
231. Nadarajah B, Parnavelas JG. Modes of neuronal migration in the developing cerebral
cortex. Nat Rev Neurosci 3: 423– 432, 2002.
232. Nakamura T, Yasuda S, Nagai H, Koinuma S, Morishita S, Goto A, Kinashi T, Wada N.
Longest neurite-specific activation of Rap1B in hippocampal neurons contributes to
polarity formation through RalA and Nore1A in addition to PI3-kinase. Genes Cells 18:
1020 –1031, 2013.
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
199. MacAskill AF, Brickley K, Stephenson FA, Kittler JT. GTPase dependent recruitment
of Grif-1 by Miro1 regulates mitochondrial trafficking in hippocampal neurons. Mol
Cell Neurosci 40: 301–312, 2009.
215. Miyoshi G, Fishell G. Dynamic FoxG1 expression coordinates the integration of multipolar pyramidal neuron precursors into the cortical plate. Neuron 74: 1045–1058,
2012.
NAMBA ET AL.
233. Nakamuta S, Funahashi Y, Namba T, Arimura N, Picciotto MR, Tokumitsu H, Soderling TR, Sakakibara A, Miyata T, Kamiguchi H, Kaibuchi K. Local application of neurotrophins specifies axons through inositol 1,4,5-trisphosphate, calcium, and Ca2⫹/
calmodulin-dependent protein kinases. Science Signaling 4: ra76, 2011.
234. Nakata T, Hirokawa N. Microtubules provide directional cues for polarized axonal
transport through interaction with kinesin motor head. J Cell Biol 162: 1045–1055,
2003.
235. Nakata T, Niwa S, Okada Y, Perez F, Hirokawa N. Preferential binding of a kinesin-1
motor to GTP-tubulin-rich microtubules underlies polarized vesicle transport. J Cell
Biol 194: 245–255, 2011.
236. Nakielny S, Cohen P, Wu J, Sturgill T. MAP kinase activator from insulin-stimulated
skeletal muscle is a protein threonine/tyrosine kinase. EMBO J 11: 2123–2129, 1992.
237. Namba T, Kibe Y, Funahashi Y, Nakamuta S, Takano T, Ueno T, Shimada A, Kozawa
S, Okamoto M, Shimoda Y, Oda K, Wada Y, Masuda T, Sakakibara A, Igarashi M,
Miyata T, Faivre-Sarrailh C, Takeuchi K, Kaibuchi K. Pioneering axons regulate neuronal polarization in the developing cerebral cortex. Neuron 81: 814 – 829, 2014.
239. Naoki H, Ishii S. Mathematical modeling of neuronal polarization during development.
Progr Mol Biol Transl Sci 123: 127–141, 2014.
240. Naoki H, Nakamuta S, Kaibuchi K, Ishii S. Flexible search for single-axon morphology
during neuronal spontaneous polarization. PLoS One 6: e19034, 2011.
241. Nguyen L, Besson A, Heng JI, Schuurmans C, Teboul L, Parras C, Philpott A, Roberts
JM, Guillemot F. p27kip1 independently promotes neuronal differentiation and migration in the cerebral cortex. Genes Dev 20: 1511–1524, 2006.
242. Nishida E, Maekawa S, Sakai H. Cofilin, a protein in porcine brain that binds to actin
filaments and inhibits their interactions with myosin and tropomyosin. Biochemistry
23: 5307–5313, 1984.
243. Nishimura T, Fukata Y, Kato K, Yamaguchi T, Matsuura Y, Kamiguchi H, Kaibuchi K.
CRMP-2 regulates polarized Numb-mediated endocytosis for axon growth. Nat Cell
Biol 5: 819 – 826, 2003.
244. Nishimura T, Kato K, Yamaguchi T, Fukata Y, Ohno S, Kaibuchi K. Role of the
PAR-3-KIF3 complex in the establishment of neuronal polarity. Nat Cell Biol 6: 328 –
334, 2004.
245. Nishimura T, Yamaguchi T, Kato K, Yoshizawa M, Nabeshima Y, Ohno S, Hoshino M,
Kaibuchi K. PAR-6-PAR-3 mediates Cdc42-induced Rac activation through the Rac
GEFs STEF/Tiam1. Nat Cell Biol 7: 270 –277, 2005.
246. Niwa S, Tanaka Y, Hirokawa N. KIF1Bbeta- and KIF1A-mediated axonal transport of
presynaptic regulator Rab3 occurs in a GTP-dependent manner through DENN/
MADD. Nat Cell Biol 10: 1269 –1279, 2008.
247. Nobes CD, Lauritzen I, Mattei MG, Paris S, Hall A, Chardin P. A new member of the
Rho family, Rnd1, promotes disassembly of actin filament structures and loss of cell
adhesion. J Cell Biol 141: 187–197, 1998.
248. Noctor SC, Flint AC, Weissman TA, Dammerman RS, Kriegstein AR. Neurons derived from radial glial cells establish radial units in neocortex. Nature 409: 714 –720,
2001.
249. Noctor SC, Martinez-Cerdeno V, Ivic L, Kriegstein AR. Cortical neurons arise in
symmetric and asymmetric division zones and migrate through specific phases. Nat
Neurosci 7: 136 –144, 2004.
250. North BJ, Marshall BL, Borra MT, Denu JM, Verdin E. The human Sir2 ortholog,
SIRT2, is an NAD⫹-dependent tubulin deacetylase. Mol Cell 11: 437– 444, 2003.
251. Ohtaka-Maruyama C, Hirai S, Miwa A, Heng JI, Shitara H, Ishii R, Taya C, Kawano H,
Kasai M, Nakajima K, Okado H. RP58 regulates the multipolar-bipolar transition of
newborn neurons in the developing cerebral cortex. Cell Reports 3: 458 – 471, 2013.
255. Ori-McKenney KM, Jan LY, Jan YN. Golgi outposts shape dendrite morphology by
functioning as sites of acentrosomal microtubule nucleation in neurons. Neuron 76:
921–930, 2012.
256. Osmani AH, Osmani SA, Morris NR. The molecular cloning and identification of a
gene product specifically required for nuclear movement in Aspergillus nidulans. J Cell
Biol 111: 543–551, 1990.
257. Ozdamar B, Bose R, Barrios-Rodiles M, Wang HR, Zhang Y, Wrana JL. Regulation of
the polarity protein Par6 by TGFbeta receptors controls epithelial cell plasticity.
Science 307: 1603–1609, 2005.
258. Pacary E, Heng J, Azzarelli R, Riou P, Castro D, Lebel-Potter M, Parras C, Bell DM,
Ridley AJ, Parsons M, Guillemot F. Proneural transcription factors regulate different
steps of cortical neuron migration through Rnd-mediated inhibition of RhoA signaling.
Neuron 69: 1069 –1084, 2011.
259. Palay SL, Sotelo C, Peters A, Orkand PM. The axon hillock and the initial segment. J
Cell Biol 38: 193–201, 1968.
260. Park KK, Liu K, Hu Y, Smith PD, Wang C, Cai B, Xu B, Connolly L, Kramvis I, Sahin M,
He Z. Promoting axon regeneration in the adult CNS by modulation of the PTEN/
mTOR pathway. Science 322: 963–966, 2008.
261. Paturle-Lafanechere L, Edde B, Denoulet P, Van Dorsselaer A, Mazarguil H, Le Caer
JP, Wehland J, Job D. Characterization of a major brain tubulin variant which cannot be
tyrosinated. Biochemistry 30: 10523–10528, 1991.
262. Paturle-Lafanechere L, Manier M, Trigault N, Pirollet F, Mazarguil 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 Sci 107: 1529 –1543, 1994.
263. Payne DM, Rossomando AJ, Martino P, Erickson AK, Her JH, Shabanowitz J, Hunt DF,
Weber MJ, Sturgill TW. Identification of the regulatory phosphorylation sites in pp42/
mitogen-activated protein kinase (MAP kinase). EMBO J 10: 885– 892, 1991.
264. Penagarikano O, Abrahams BS, Herman EI, Winden KD, Gdalyahu A, Dong H, Sonnenblick LI, Gruver R, Almajano J, Bragin A, Golshani P, Trachtenberg JT, Peles E,
Geschwind DH. Absence of CNTNAP2 leads to epilepsy, neuronal migration abnormalities, and core autism-related deficits. Cell 147: 235–246, 2011.
265. Peretti D, Peris L, Rosso S, Quiroga S, Caceres A. Evidence for the involvement of
KIF4 in the anterograde transport of L1-containing vesicles. J Cell Biol 149: 141–152,
2000.
266. Peris L, Wagenbach M, Lafanechere L, Brocard J, Moore AT, Kozielski F, Job D,
Wordeman L, Andrieux A. Motor-dependent microtubule disassembly driven by tubulin tyrosination. J Cell Biol 185: 1159 –1166, 2009.
267. Pinyol R, Haeckel A, Ritter A, Qualmann B, Kessels MM. Regulation of N-WASP and
the Arp2/3 complex by Abp1 controls neuronal morphology. PLoS One 2: e400, 2007.
268. Poirier K, Keays DA, Francis F, Saillour Y, Bahi N, Manouvrier S, Fallet-Bianco C,
Pasquier L, Toutain A, Tuy FP, Bienvenu T, Joriot S, Odent S, Ville D, Desguerre I,
Goldenberg A, Moutard ML, Fryns JP, van Esch H, Harvey RJ, Siebold C, Flint J,
Beldjord C, Chelly J. Large spectrum of lissencephaly and pachygyria phenotypes
resulting from de novo missense mutations in tubulin alpha 1A (TUBA1A). Human
Mutat 28: 1055–1064, 2007.
269. Prasad BC, Clark SG. Wnt signaling establishes anteroposterior neuronal polarity and
requires retromer in C. elegans. Development 133: 1757–1766, 2006.
270. Prinetti A, Prioni S, Chigorno V, Karagogeos D, Tettamanti G, Sonnino S. Immunoseparation of sphingolipid-enriched membrane domains enriched in Src family protein tyrosine kinases and in the neuronal adhesion molecule TAG-1 by anti-GD3
ganglioside monoclonal antibody. J Neurochem 78: 1162–1167, 2001.
271. Prota AE, Magiera MM, Kuijpers M, Bargsten K, Frey D, Wieser M, Jaussi R, Hoogenraad CC, Kammerer RA, Janke C, Steinmetz MO. Structural basis of tubulin tyrosination by tubulin tyrosine ligase. J Cell Biol 200: 259 –270, 2013.
252. Oinuma I, Katoh H, Negishi M. R-Ras controls axon specification upstream of glycogen synthase kinase-3beta through integrin-linked kinase. J Biol Chem 282: 303–318,
2007.
272. Qin J, Wu C. ILK: a pseudokinase in the center stage of cell-matrix adhesion and
signaling. Curr Opin Cell Biol 24: 607– 613, 2012.
253. Okada Y, Yamazaki H, Sekine-Aizawa Y, Hirokawa N. The neuron-specific kinesin
superfamily protein KIF1A is a unique monomeric motor for anterograde axonal
transport of synaptic vesicle precursors. Cell 81: 769 –780, 1995.
273. Qiu RG, Abo A, Steven Martin G. A human homolog of the C. elegans polarity determinant Par-6 links Rac and Cdc42 to PKCzeta signaling and cell transformation. Curr
Biol 10: 697–707, 2000.
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
1021
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
238. Namba T, Nakamuta S, Funahashi Y, Kaibuchi K. The role of selective transport in
neuronal polarization. Dev Neurobiol 71: 445– 457, 2011.
254. Oliva AA Jr, Atkins CM, Copenagle L, Banker GA. Activated c-Jun N-terminal kinase
is required for axon formation. J Neurosci 26: 9462–9470, 2006.
NEURONAL POLARITY SIGNALING
274. Qureshi IA, Gokhan S, Mehler MF. REST and CoREST are transcriptional and epigenetic regulators of seminal neural fate decisions. Cell Cycle 9: 4477– 4486, 2010.
297. Schroer TA, Sheetz MP. Two activators of microtubule-based vesicle transport. J Cell
Biol 115: 1309 –1318, 1991.
275. Randlett O, Norden C, Harris WA. The vertebrate retina: a model for neuronal
polarization in vivo. Dev Neurobiol 71: 567–583, 2011.
298. Schwamborn JC, Puschel AW. The sequential activity of the GTPases Rap1B and
Cdc42 determines neuronal polarity. Nat Neurosci 7: 923–929, 2004.
276. Randlett O, Poggi L, Zolessi FR, Harris WA. The oriented emergence of axons from
retinal ganglion cells is directed by laminin contact in vivo. Neuron 70: 266 –280, 2011.
299. Sengottuvel V, Leibinger M, Pfreimer M, Andreadaki A, Fischer D. Taxol facilitates
axon regeneration in the mature CNS. J Neurosci 31: 2688 –2699, 2011.
277. Raper J, Mason C. Cellular strategies of axonal pathfinding. Cold Spring Harb Perspect
Biol 2: a001933, 2010.
300. Setou M, Nakagawa T, Seog DH, Hirokawa N. Kinesin superfamily motor protein
KIF17 and mLin-10 in NMDA receptor-containing vesicle transport. Science 288:
1796 –1802, 2000.
278. Rasband MN. The axon initial segment and the maintenance of neuronal polarity. Nat
Rev Neurosci 11: 552–562, 2010.
279. Raybin D, Flavin M. Enzyme which specifically adds tyrosine to the alpha chain of
tubulin. Biochemistry 16: 2189 –2194, 1977.
280. Reed NA, Cai D, Blasius TL, Jih GT, Meyhofer E, Gaertig J, Verhey KJ. Microtubule
acetylation promotes kinesin-1 binding and transport. Curr Biol 16: 2166 –2172, 2006.
282. Reiner O, Carrozzo R, Shen Y, Wehnert M, Faustinella F, Dobyns WB, Caskey CT,
Ledbetter DH. Isolation of a Miller-Dieker lissencephaly gene containing G protein
beta-subunit-like repeats. Nature 364: 717–721, 1993.
283. Reiner O, Sapir T. Polarity regulation in migrating neurons in the cortex. Mol Neurobiol
40: 1–14, 2009.
284. Sahab ZJ, Hall MD, Me Sung Y, Dakshanamurthy S, Ji Y, Kumar D, Byers SW. Tumor
suppressor RARRES1 interacts with cytoplasmic carboxypeptidase AGBL2 to regulate
the alpha-tubulin tyrosination cycle. Cancer Res 71: 1219 –1228, 2011.
302. Shelly M, Cancedda L, Heilshorn S, Sumbre G, Poo MM. LKB1/STRAD promotes axon
initiation during neuronal polarization. Cell 129: 565–577, 2007.
303. Shelly M, Cancedda L, Lim BK, Popescu AT, Cheng PL, Gao H, Poo MM.
Semaphorin3A regulates neuronal polarization by suppressing axon formation and
promoting dendrite growth. Neuron 71: 433– 446, 2011.
304. Shelly M, Lim BK, Cancedda L, Heilshorn SC, Gao H, Poo MM. Local and long-range
reciprocal regulation of cAMP and cGMP in axon/dendrite formation. Science 327:
547–552, 2010.
305. Shi SH, Jan LY, Jan YN. Hippocampal neuronal polarity specified by spatially localized
mPar3/mPar6 and PI 3-kinase activity. Cell 112: 63–75, 2003.
306. Shida T, Cueva JG, Xu Z, Goodman MB, Nachury MV. The major alpha-tubulin
K40 acetyltransferase alphaTAT1 promotes rapid ciliogenesis and efficient
mechanosensation. Proc Natl Acad Sci USA 107: 21517–21522, 2010.
285. Saito T, Nakatsuji N. Efficient gene transfer into the embryonic mouse brain using in
vivo electroporation. Dev Biol 240: 237–246, 2001.
307. Shimada T, Toriyama M, Uemura K, Kamiguchi H, Sugiura T, Watanabe N, Inagaki N.
Shootin1 interacts with actin retrograde flow and L1-CAM to promote axon outgrowth. J Cell Biol 181: 817– 829, 2008.
286. Sakakibara A, Ando R, Sapir T, Tanaka T. Microtubule dynamics in neuronal morphogenesis. Open Biol 3: 130061, 2013.
308. Shimoda Y, Watanabe K. Contactins: emerging key roles in the development and
function of the nervous system. Cell Adhesion Migration 3: 64 –70, 2009.
287. Sakakibara A, Sato T, Ando R, Noguchi N, Masaoka M, Miyata T. Dynamics of
centrosome translocation and microtubule organization in neocortical neurons
during distinct modes of polarization. Cereb Cortex 24: 1301–1310,
2014.
309. Shinoda T, Taya S, Tsuboi D, Hikita T, Matsuzawa R, Kuroda S, Iwamatsu A, Kaibuchi
K. DISC1 regulates neurotrophin-induced axon elongation via interaction with Grb2.
J Neurosci 27: 4 –14, 2007.
288. Sakane A, Manabe S, Ishizaki H, Tanaka-Okamoto M, Kiyokage E, Toida K, Yoshida T,
Miyoshi J, Kamiya H, Takai Y, Sasaki T. Rab3 GTPase-activating protein regulates
synaptic transmission and plasticity through the inactivation of Rab3. Proc Natl Acad Sci
USA 103: 10029 –10034, 2006.
289. Sampo B, Kaech S, Kunz S, Banker G. Two distinct mechanisms target membrane
proteins to the axonal surface. Neuron 37: 611– 624, 2003.
290. Sapir T, Levy T, Sakakibara A, Rabinkov A, Miyata T, Reiner O. Shootin1 acts in
concert with KIF20B to promote polarization of migrating neurons. J Neurosci 33:
11932–11948, 2013.
291. Sapir T, Sapoznik S, Levy T, Finkelshtein D, Shmueli A, Timm T, Mandelkow EM,
Reiner O. Accurate balance of the polarity kinase MARK2/Par-1 is required for proper
cortical neuronal migration. J Neurosci 28: 5710 –5720, 2008.
292. Sapir T, Shmueli A, Levy T, Timm T, Elbaum M, Mandelkow EM, Reiner O. Antagonistic effects of doublecortin and MARK2/Par-1 in the developing cerebral cortex. J
Neurosci 28: 13008 –13013, 2008.
310. Shu T, Ayala R, Nguyen MD, Xie Z, Gleeson JG, Tsai LH. Ndel1 operates in a common
pathway with LIS1 and cytoplasmic dynein to regulate cortical neuronal positioning.
Neuron 44: 263–277, 2004.
311. Simons M, Mlodzik M. Planar cell polarity signaling: from fly development to human
disease. Annu Rev Genet 42: 517–540, 2008.
312. Smith DS, Niethammer M, Ayala R, Zhou Y, Gambello MJ, Wynshaw-Boris A, Tsai LH.
Regulation of cytoplasmic dynein behaviour and microtubule organization by mammalian Lis1. Nat Cell Biol 2: 767–775, 2000.
313. Smith MJ, Pozo K, Brickley K, Stephenson FA. Mapping the GRIF-1 binding domain of
the kinesin, KIF5C, substantiates a role for GRIF-1 as an adaptor protein in the
anterograde trafficking of cargoes. J Biol Chem 281: 27216 –27228, 2006.
314. Sobotzik JM, Sie JM, Politi C, Del Turco D, Bennett V, Deller T, Schultz C. AnkyrinG
is required to maintain axo-dendritic polarity in vivo. Proc Natl Acad Sci USA 106:
17564 –17569, 2009.
315. Song AH, Wang D, Chen G, Li Y, Luo J, Duan S, Poo MM. A selective filter for
cytoplasmic transport at the axon initial segment. Cell 136: 1148 –1160, 2009.
293. Schaar BT, Kinoshita K, McConnell SK. Doublecortin microtubule affinity is regulated
by a balance of kinase and phosphatase activity at the leading edge of migrating
neurons. Neuron 41: 203–213, 2004.
316. Song Y, Kirkpatrick LL, Schilling AB, Helseth DL, Chabot N, Keillor JW, Johnson GV,
Brady ST. Transglutaminase and polyamination of tubulin: posttranslational modification for stabilizing axonal microtubules. Neuron 78: 109 –123, 2013.
294. Schaar BT, McConnell SK. Cytoskeletal coordination during neuronal migration. Proc
Natl Acad Sci USA 102: 13652–13657, 2005.
317. St Johnston D, Ahringer J. Cell polarity in eggs and epithelia: parallels and diversity. Cell
141: 757–774, 2010.
295. Schlager MA, Hoogenraad CC. Basic mechanisms for recognition and transport of
synaptic cargos. Mol Brain 2: 25, 2009.
318. Stenmark H. Rab GTPases as coordinators of vesicle traffic. Nat Rev Mol Cell Biol 10:
513–525, 2009.
296. Schroder HC, Wehland J, Weber K. Purification of brain tubulin-tyrosine ligase by
biochemical and immunological methods. J Cell Biol 100: 276 –281, 1985.
319. Stepanova T, Slemmer J, Hoogenraad CC, Lansbergen G, Dortland B, De Zeeuw CI,
Grosveld F, van Cappellen G, Akhmanova A, Galjart N. Visualization of microtubule
1022
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
281. Reichardt LF. Neurotrophin-regulated signalling pathways. Philos Trans R Soc Lond B
Biol Sci 361: 1545–1564, 2006.
301. Setou M, Seog DH, Tanaka Y, Kanai Y, Takei Y, Kawagishi M, Hirokawa N. Glutamatereceptor-interacting protein GRIP1 directly steers kinesin to dendrites. Nature 417:
83– 87, 2002.
NAMBA ET AL.
growth in cultured neurons via the use of EB3-GFP (end-binding protein 3-green
fluorescent protein). J Neurosci 23: 2655–2664, 2003.
341. Tissir F, Goffinet AM. Reelin and brain development. Nat Rev Neurosci 4: 496 –505,
2003.
320. Stiess M, Maghelli N, Kapitein LC, Gomis-Ruth S, Wilsch-Brauninger M, Hoogenraad
CC, Tolic-Norrelykke IM, Bradke F. Axon extension occurs independently of centrosomal microtubule nucleation. Science 327: 704 –707, 2010.
342. Tonosaki M, Itoh K, Umekage M, Kishimoto T, Yaoi T, Lemmon VP, Fushiki S. L1cam
is crucial for cell locomotion and terminal translocation of the soma in radial migration
during murine corticogenesis. PLoS One 9: e86186, 2014.
321. Stoorvogel W. Functional transfer of microRNA by exosomes. Blood 119: 646 – 648,
2012.
343. Toriyama M, Kozawa S, Sakumura Y, Inagaki N. Conversion of a signal into forces for
axon outgrowth through Pak1-mediated shootin1 phosphorylation. Curr Biol 23: 529 –
534, 2013.
322. Stowers RS, Megeath LJ, Gorska-Andrzejak J, Meinertzhagen IA, Schwarz TL. Axonal
transport of mitochondria to synapses depends on milton, a novel Drosophila protein.
Neuron 36: 1063–1077, 2002.
323. Sugioka K, Sawa H. Formation and functions of asymmetric microtubule organization
in polarized cells. Curr Opin Cell Biol 24: 517–525, 2012.
324. Sun Y, Fei T, Yang T, Zhang F, Chen YG, Li H, Xu Z. The suppression of CRMP2
expression by bone morphogenetic protein (BMP)-SMAD gradient signaling controls
multiple stages of neuronal development. J Biol Chem 285: 39039 –39050, 2010.
326. Svitkina TM, Verkhovsky AB, McQuade KM, Borisy GG. Analysis of the actin-myosin
II system in fish epidermal keratocytes: mechanism of cell body translocation. J Cell
Biol 139: 397– 415, 1997.
327. Szu-Yu Ho T, Rasband MN. Maintenance of neuronal polarity. Dev Neurobiol 71:
474 – 482, 2011.
328. Tabata H, Nakajima K. Efficient in utero gene transfer system to the developing
mouse brain using electroporation: visualization of neuronal migration in the developing cortex. Neuroscience 103: 865– 872, 2001.
329. Tabata H, Nakajima K. Multipolar migration: the third mode of radial neuronal migration in the developing cerebral cortex. J Neurosci 23: 9996 –10001, 2003.
330. Tahirovic S, Hellal F, Neukirchen D, Hindges R, Garvalov BK, Flynn KC, Stradal TE,
Chrostek-Grashoff A, Brakebusch C, Bradke F. Rac1 regulates neuronal polarization
through the WAVE complex. J Neurosci 30: 6930 – 6943, 2010.
345. Toriyama M, Shimada T, Kim KB, Mitsuba M, Nomura E, Katsuta K, Sakumura Y,
Roepstorff P, Inagaki N. Shootin1: a protein involved in the organization of an asymmetric signal for neuronal polarization. J Cell Biol 175: 147–157, 2006.
346. Tsai JW, Bremner KH, Vallee RB. Dual subcellular roles for LIS1 and dynein in radial
neuronal migration in live brain tissue. Nat Neurosci 10: 970 –979, 2007.
347. Tsai LH, Gleeson JG. Nucleokinesis in neuronal migration. Neuron 46: 383–388,
2005.
348. Tsuboi D, Hikita T, Qadota H, Amano M, Kaibuchi K. Regulatory machinery of
UNC-33 Ce-CRMP localization in neurites during neuronal development in Caenorhabditis elegans. J Neurochem 95: 1629 –1641, 2005.
349. Turing AM. The chemical basis of morphogenesis. 1953 Bull Math Biol 52: 119 –197,
1990.
350. Twelvetrees AE, Yuen EY, Arancibia-Carcamo IL, MacAskill AF, Rostaing P, Lumb MJ,
Humbert S, Triller A, Saudou F, Yan Z, Kittler JT. Delivery of GABAARs to synapses is
mediated by HAP1-KIF5 and disrupted by mutant huntingtin. Neuron 65: 53– 65,
2010.
351. Uboha NV, Flajolet M, Nairn AC, Picciotto MR. A calcium- and calmodulin-dependent
kinase Ialpha/microtubule affinity regulating kinase 2 signaling cascade mediates calcium-dependent neurite outgrowth. J Neurosci 27: 4413– 4423, 2007.
331. Takai Y, Sasaki T, Matozaki T. Small GTP-binding proteins. Physiol Rev 81: 153–208,
2001.
352. Umeshima H, Hirano T, Kengaku M. Microtubule-based nuclear movement occurs
independently of centrosome positioning in migrating neurons. Proc Natl Acad Sci USA
104: 16182–16187, 2007.
332. Takatalo M, Jarvinen E, Laitinen S, Thesleff I, Ronnholm R. Expression of the novel
Golgi protein GoPro49 is developmentally regulated during mesenchymal differentiation. Dev Dyn 237: 2243–2255, 2008.
353. Vallee RB, McKenney RJ, Ori-McKenney KM. Multiple modes of cytoplasmic dynein
regulation. Nat Cell Biol 14: 224 –230, 2012.
333. Tamamaki N, Nakamura K, Okamoto K, Kaneko T. Radial glia is a progenitor of
neocortical neurons in the developing cerebral cortex. Neurosci Res 41: 51– 60, 2001.
354. Verhey KJ, Meyer D, Deehan R, Blenis J, Schnapp BJ, Rapoport TA, Margolis B. Cargo
of kinesin identified as JIP scaffolding proteins and associated signaling molecules. J Cell
Biol 152: 959 –970, 2001.
334. Tanaka N, Meng W, Nagae S, Takeichi M. Nezha/CAMSAP3 and CAMSAP2 cooperate in epithelial-specific organization of noncentrosomal microtubules. Proc Natl Acad
Sci USA 109: 20029 –20034, 2012.
355. Vojtek AB, Hollenberg SM, Cooper JA. Mammalian Ras interacts directly with the
serine/threonine kinase Raf. Cell 74: 205–214, 1993.
335. Tanaka T, Serneo FF, Higgins C, Gambello MJ, Wynshaw-Boris A, Gleeson JG. Lis1
and doublecortin function with dynein to mediate coupling of the nucleus to the
centrosome in neuronal migration. J Cell Biol 165: 709 –721, 2004.
356. Volvert ML, Prevot PP, Close P, Laguesse S, Pirotte S, Hemphill J, Rogister F, Kruzy N,
Sacheli R, Moonen G, Deiters A, Merkenschlager M, Chariot A, Malgrange B, Godin
JD, Nguyen L. MicroRNA targeting of CoREST controls polarization of migrating
cortical neurons. Cell Reports 7: 1168 –1183, 2014.
336. Tapia M, Wandosell F, Garrido JJ. Impaired function of HDAC6 slows down axonal
growth and interferes with axon initial segment development. PLoS One 5: e12908,
2010.
337. Tararuk T, Ostman N, Li W, Bjorkblom B, Padzik A, Zdrojewska J, Hongisto V,
Herdegen T, Konopka W, Courtney MJ, Coffey ET. JNK1 phosphorylation of SCG10
determines microtubule dynamics and axodendritic length. J Cell Biol 173: 265–277,
2006.
338. Taverna E, Gotz M, Huttner WB. The cell biology of neurogenesis: toward an understanding of the development and evolution of the neocortex. Annu Rev Cell Dev Biol 30:
465–502, 2014.
339. Taya S, Shinoda T, Tsuboi D, Asaki J, Nagai K, Hikita T, Kuroda S, Kuroda K, Shimizu
M, Hirotsune S, Iwamatsu A, Kaibuchi K. DISC1 regulates the transport of the NUDEL/LIS1/14-3-3epsilon complex through kinesin-1. J Neurosci 27: 15–26, 2007.
340. Terada S, Kinjo M, Hirokawa N. Oligomeric tubulin in large transporting complex is
transported via kinesin in squid giant axons. Cell 103: 141–155, 2000.
357. Voss AK, Britto JM, Dixon MP, Sheikh BN, Collin C, Tan SS, Thomas T. C3G regulates
cortical neuron migration, preplate splitting and radial glial cell attachment. Development 135: 2139 –2149, 2008.
358. Wang X, Destrument A, Tournier C. Physiological roles of MKK4 and MKK7: insights
from animal models. Biochim Biophys Acta 1773: 1349 –1357, 2007.
359. Wang X, Schwarz TL. The mechanism of Ca2⫹-dependent regulation of kinesinmediated mitochondrial motility. Cell 136: 163–174, 2009.
360. Waterman-Storer CM, Karki SB, Kuznetsov SA, Tabb JS, Weiss DG, Langford GM,
Holzbaur EL. The interaction between cytoplasmic dynein and dynactin is required
for fast axonal transport. Proc Natl Acad Sci USA 94: 12180 –12185, 1997.
361. Watts JL, Etemad-Moghadam B, Guo S, Boyd L, Draper BW, Mello CC, Priess JR,
Kemphues KJ. par-6, a gene involved in the establishment of asymmetry in early C.
elegans embryos, mediates the asymmetric localization of PAR-3. Development 122:
3133–3140, 1996.
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
1023
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
325. Suzuki A, Ohno S. The PAR-aPKC system: lessons in polarity. J Cell Sci 119: 979 –987,
2006.
344. Toriyama M, Sakumura Y, Shimada T, Ishii S, Inagaki N. A diffusion-based neurite
length-sensing mechanism involved in neuronal symmetry breaking. Mol Syst Biol 6:
394, 2010.
NEURONAL POLARITY SIGNALING
362. Wayman GA, Kaech S, Grant WF, Davare M, Impey S, Tokumitsu H, Nozaki N,
Banker G, Soderling TR. Regulation of axonal extension and growth cone motility by
calmodulin-dependent protein kinase I. J Neurosci 24: 3786 –3794, 2004.
363. Webster DR, Borisy GG. Microtubules are acetylated in domains that turn over
slowly. J Cell Sci 92: 57– 65, 1989.
364. Wennerberg K, Forget MA, Ellerbroek SM, Arthur WT, Burridge K, Settleman J, Der
CJ, Hansen SH. Rnd proteins function as RhoA antagonists by activating p190
RhoGAP. Curr Biol 13: 1106 –1115, 2003.
365. Westerlund N, Zdrojewska J, Padzik A, Komulainen E, Bjorkblom B, Rannikko E,
Tararuk T, Garcia-Frigola C, Sandholm J, Nguyen L, Kallunki T, Courtney MJ, Coffey
ET. Phosphorylation of SCG10/stathmin-2 determines multipolar stage exit and neuronal migration rate. Nat Neurosci 14: 305–313, 2011.
366. Whitmarsh AJ, Kuan CY, Kennedy NJ, Kelkar N, Haydar TF, Mordes JP, Appel M,
Rossini AA, Jones SN, Flavell RA, Rakic P, Davis RJ. Requirement of the JIP1 scaffold
protein for stress-induced JNK activation. Genes Dev 15: 2421–2432, 2001.
373. Yamaguchi K, Tanaka M, Mizoguchi A, Hirata Y, Ishizaki H, Kaneko K, Miyoshi J, Takai
Y. A GDP/GTP exchange protein for the Rab3 small G protein family up-regulates a
postdocking step of synaptic exocytosis in central synapses. Proc Natl Acad Sci USA 99:
14536 –14541, 2002.
374. Yamashita N, Ohshima T, Nakamura F, Kolattukudy P, Honnorat J, Mikoshiba K,
Goshima Y. Phosphorylation of CRMP2 (collapsin response mediator protein 2) is involved in proper dendritic field organization. J Neurosci 32: 1360 –1365, 2012.
375. Yau KW, van Beuningen SF, Cunha-Ferreira I, Cloin BM, van Battum EY, Will L,
Schatzle P, Tas RP, van Krugten J, Katrukha EA, Jiang K, Wulf PS, Mikhaylova M,
Harterink M, Pasterkamp RJ, Akhmanova A, Kapitein LC, Hoogenraad CC. Microtubule minus-end binding protein CAMSAP2 controls axon specification and dendrite
development. Neuron 82: 1058 –1073, 2014.
376. Yi JJ, Barnes AP, Hand R, Polleux F, Ehlers MD. TGF-beta signaling specifies axons
during brain development. Cell 142: 144 –157, 2010.
377. Yoshimura T, Arimura N, Kawano Y, Kawabata S, Wang S, Kaibuchi K. Ras regulates
neuronal polarity via the PI3-kinase/Akt/GSK-3beta/CRMP-2 pathway. Biochem Biophys Res Commun 340: 62– 68, 2006.
368. Witte H, Neukirchen D, Bradke F. Microtubule stabilization specifies initial neuronal
polarization. J Cell Biol 180: 619 – 632, 2008.
378. Yoshimura T, Kawano Y, Arimura N, Kawabata S, Kikuchi A, Kaibuchi K. GSK-3beta
regulates phosphorylation of CRMP-2 and neuronal polarity. Cell 120: 137–149, 2005.
369. Wu SX, Goebbels S, Nakamura K, Kometani K, Minato N, Kaneko T, Nave KA,
Tamamaki N. Pyramidal neurons of upper cortical layers generated by NEX-positive
progenitor cells in the subventricular zone. Proc Natl Acad Sci USA 102: 17172–17177,
2005.
379. Yoshimura Y, Terabayashi T, Miki H. Par1b/MARK2 phosphorylates kinesin-like motor
protein GAKIN/KIF13B to regulate axon formation. Mol Cell Biol 30: 2206 –2219, 2010.
370. Xia CH, Roberts EA, Her LS, Liu X, Williams DS, Cleveland DW, Goldstein LS.
Abnormal neurofilament transport caused by targeted disruption of neuronal kinesin
heavy chain KIF5A. J Cell Biol 161: 55– 66, 2003.
371. Yabe JT, Pimenta A, Shea TB. Kinesin-mediated transport of neurofilament protein
oligomers in growing axons. J Cell Sci 112: 3799 –3814, 1999.
372. Yamada M, Toba S, Takitoh T, Yoshida Y, Mori D, Nakamura T, Iwane AH, Yanagida
T, Imai H, Yu-Lee LY, Schroer T, Wynshaw-Boris A, Hirotsune S. mNUDC is required
for plus-end-directed transport of cytoplasmic dynein and dynactins by kinesin-1.
EMBO J 29: 517–531, 2010.
1024
380. Yoshizawa M, Kawauchi T, Sone M, Nishimura YV, Terao M, Chihama K, Nabeshima
Y, Hoshino M. Involvement of a Rac activator,P-Rex1, in neurotrophin-derived signaling and neuronal migration. J Neurosci 25: 4406 – 4419, 2005.
381. Zerial M, McBride H. Rab proteins as membrane organizers. Nat Rev Mol Cell Biol 2:
107–117, 2001.
382. Zhao C, Takita J, Tanaka Y, Setou M, Nakagawa T, Takeda S, Yang HW, Terada S, Nakata
T, Takei Y, Saito M, Tsuji S, Hayashi Y, Hirokawa N. Charcot-Marie-Tooth disease type
2A caused by mutation in a microtubule motor KIF1Bbeta. Cell 105: 587–597, 2001.
383. Zolessi FR, Poggi L, Wilkinson CJ, Chien CB, Harris WA. Polarization and orientation
of retinal ganglion cells in vivo. Neural Dev 1: 2, 2006.
Physiol Rev • VOL 95 • JULY 2015 • www.prv.org
Downloaded from http://physrev.physiology.org/ by 10.220.33.2 on May 3, 2017
367. Witte H, Bradke F. The role of the cytoskeleton during neuronal polarization. Curr
Opin Neurobiol 18: 479 – 487, 2008.